DNA Nanotechnology: Methods and Protocols (Methods in Molecular Biology Vol 749)

Methods

in

Molecular Biology™

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



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

DNA Nanotechnology Methods and Protocols Edited by

Giampaolo Zuccheri and Bruno Samorì Department of Biochemistry, University of Bologna, Bologna, Italy

Editors Giampaolo Zuccheri, Ph.D. Department of Biochemistry University of Bologna Bologna, Italy [email protected]

Bruno Samorì Department of Biochemistry University of Bologna Bologna, Italy [email protected]

ISSN 1064-3745 e-ISSN 1940-6029 ISBN 978-1-61779-141-3 e-ISBN 978-1-61779-142-0 DOI 10.1007/978-1-61779-142-0 Springer New York Heidelberg London Dordrecht Library of Congress Control Number: 2011929163 © Springer Science+Business Media, LLC 2011 All rights reserved. This work may not be translated or copied in whole or in part without the written permission of the publisher (Humana Press, c/o Springer Science+Business Media, LLC, 233 Spring Street, New York, NY 10013, USA), except for brief excerpts in connection with reviews or scholarly analysis. Use in connection with any form of information storage and retrieval, electronic adaptation, computer software, or by similar or ­dissimilar methodology now known or hereafter developed is forbidden. The use in this publication of trade names, trademarks, service marks, and similar terms, even if they are not identified as such, is not to be taken as an expression of opinion as to whether or not they are subject to proprietary rights. Printed on acid-free paper Humana Press is part of Springer Science+Business Media (www.springer.com)

Preface Giorgio Vasari, a painter, architect, and art historian during the Italian Renaissance, is credited with coining the expression “andare a bottega,” (“attending the studio”) referring to the internship that the apprentice would complete in the master’s studio in order to learn what could be uniquely transmitted in person and in that particular environment and that could then lead to making a unique artist of the apprentice. Nowadays, this same concept holds true in science, and despite the many opportunities for communication and “virtual presence”, the real physical permanence in a lab is still the best way for a scientist to learn a technique or a protocol, or a way of thinking. A book of protocols, such as this, humbly proposes itself as the second-best option. Not quite the same as being in person in a lab and witnessing the experts’ execution of a protocol, it still holds many more details and hints than the usually brief methods section found in research papers. This book of protocols for DNA nanotechnology was composed with this concept in mind: prolonging the tradition of Methods in Molecular Biology, it tries to simplify researchers’ lives when they are putting in practice protocols whose results they have learnt in scientific journals. DNA is playing a quite important and dual role in nanotechnology. First, its properties can nowadays be studied with unprecedented detail, thanks to the new instrumental nano(bio)technologies and new insight is being gathered on the biological behavior and function of DNA thanks to new instrumentation, smart experimental design, and protocols. Second, the DNA molecule can be decontextualized and “simply” used as a copolymer with designed interaction rules. The Watson–Crick pairing code can be harnessed towards implementing the most complicated and elegant molecular self-assembly reported to date. After Ned Seeman’s contribution, elegantly complicated branched structures can be braided and joined towards building nano-objects of practically any desired form. DNA nanotechnology is somewhat like watching professional tennis players: everything seems so simple, but then you set foot on the court and realize how difficult it is to hit a nice shot. When you see the structural perfection of a self-assembling DNA nanoobject, such as a DNA origami, you marvel at how smart DNA is as a molecule and wonder how many different constructs you could design and realize. Among the others, this book tries to show the procedures to follow in order to repeat some of the methods that lead to such constructs, or to the mastering of the characterization techniques used to study them. Many details and procedures are the fruit of the blending of artistry, science, and patience, which are often unseen in a journal paper, but that could be what makes the difference between a winning shot and hitting the net. Many research groups share their expertise with the readers in this book. For the sake of conciseness, we here mention the group leaders, while it is truly from the daily work of a complete team that the details of a protocol can be worked out. The chapters of this book can be roughly divided into two parts: some deal with the methods of preparing the nanostructures, from the rationale of the operations to the techniques for their handling; some other chapters deal more directly with advanced instrumental techniques that can manipulate and characterize molecules and nanostructures. As part of the first group, Roberto Corradini introduces the reader to the methods and choices for taming helix chirality, Alexander Kotlyar, Wolfgang Fritzsche, Naoki Sugimoto, and James Vesenka

v

vi

Preface

share their different methods in growing, characterizing, and modifying nanowires based on G tetraplexes; Hao Yan and Friedrich Simmel teach all the basics for implementing the self-assembly of branched DNA nanostructures, and then characterizing the assembly. Hanadi Sleiman tells about hybrid metal–DNA nanostructures with controlled geometry. Frank Bier shows the use of rolling circle amplification to make repetitive DNA nanostructures, while, moving closer to technological use of DNA, Arianna Filoramo instructs on how to metalize double-stranded DNA and Andrew Houlton reports on the protocol to grow DNA oligonucleotides on silicon. Also with an eye to the applicative side, Yamuna Krishnan instructs on how to insert and use DNA nanostructures inside living cells. On the instrument side, Ciro Cecconi and Mark Williams introduce the readers to methods for the use of optical tweezers, focusing mainly on the preparation of the ideal molecular construct and on the instrument and its handling, respectively. John van Noort and Sanford Leuba give us protocols on how to obtain sound data from single-molecule FRET and apply it to study the structure of chromatin. Claudio Rivetti teaches the reader how to extract quantitative data from AFM of DNA and its complexes, while Matteo Castronovo instructs on the subtleties of using the AFM as a nanolithography tool on self-assembled monolayers; Jussi Toppari dwelves on the very interesting use of dielectrophoresis as a method to manipulate and confine DNA, while Matteo Palma and Jennifer Cha explain methods for confining on surfaces DNA and those very same types of DNA nanostructures that other chapters tell the reader how to assemble. Aleksei Aksimientev shows the methods for modeling nanopores for implementing DNA translocation, a technique bound to find many applications in the near future. We hope this book will help ignite interest and spur activity in this young research field, expanding our family of enthusiastic followers and practitioners. There are certainly still many chapters to be written on this subject, simply because so much is happening in the labs at this very moment. There will certainly be room for the mainstreaming of protocols on the use of DNA analogues (starting with the marvelous RNA, of course), for the design and preparation of fully 3D architectures, for the development of routes towards functional DNA nanostructures, which will lead to applications. DNA nanostructures can be “re-inserted” in their original biological context, as microorganisms can be convinced to replicate nanostructures or even code them. And eventually, applications will require massive amounts of the nanostructures to be produced and to be manipulated automatically, possibly with a precision and output rate similar to that of the assembly of microelectronics circuitry nowadays. Our personal wish is that the next chapters will be written by some of our readers. Bologna, Italy Bologna, Italy

Giampaolo Zuccheri Bruno Samorì

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

v ix

  1 Synthesis and Characterization of Self-Assembled DNA Nanostructures . . . . . . . . Chenxiang Lin, Yonggang Ke, Rahul Chhabra, Jaswinder Sharma, Yan Liu, and Hao Yan   2 Protocols for Self-Assembly and Imaging of DNA Nanostructures . . . . . . . . . . . . Thomas L. Sobey and Friedrich C. Simmel   3 Self-Assembly of Metal-DNA Triangles and DNA Nanotubes with Synthetic Junctions . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Hua Yang, Pik Kwan Lo, Christopher K. McLaughlin, Graham D. Hamblin, Faisal A. Aldaye, and Hanadi F. Sleiman   4 DNA-Templated Pd Conductive Metallic Nanowires . . . . . . . . . . . . . . . . . . . . . . Khoa Nguyen, Stephane Campidelli, and Arianna Filoramo   5 A Method to Map Spatiotemporal pH Changes Inside Living Cells Using a pH-Triggered DNA Nanoswitch . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Souvik Modi and Yamuna Krishnan   6 Control of Helical Handedness in DNA and PNA Nanostructures . . . . . . . . . . . . Roberto Corradini, Tullia Tedeschi, Stefano Sforza, Mark M. Green, and Rosangela Marchelli   7 G-Quartet, G-Quadruplex, and G-Wire Regulated by Chemical Stimuli . . . . . . . . Daisuke Miyoshi and Naoki Sugimoto   8 Preparation and Atomic Force Microscopy of Quadruplex DNA . . . . . . . . . . . . . James Vesenka   9 Synthesis of Long DNA-Based Nanowires . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Alexander Kotlyar 10 G-Wire Synthesis and Modification with Gold Nanoparticle . . . . . . . . . . . . . . . . . Christian Leiterer, Andrea Csaki, and Wolfgang Fritzsche 11 Preparation of DNA Nanostructures with Repetitive Binding Motifs by Rolling Circle Amplification . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Edda Reiß, Ralph Hölzel, and Frank F. Bier 12 Controlled Confinement of DNA at the Nanoscale: Nanofabrication and Surface Bio-Functionalization . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Matteo Palma, Justin J. Abramson, Alon A. Gorodetsky, Colin Nuckolls, Michael P. Sheetz, Shalom J. Wind, and James Hone 13 Templated Assembly of DNA Origami Gold Nanoparticle Arrays on Lithographically Patterned Surfaces . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Albert M. Hung and Jennifer N. Cha 14 DNA-Modified Single Crystal and Nanoporous Silicon . . . . . . . . . . . . . . . . . . . . Andrew Houlton, Bernard A. Connolly, Andrew R. Pike, and Benjamin R. Horrocks

1

vii

13

33

49

61 79

93 105 115 141

151

169

187 199

viii

Contents

15 The Atomic Force Microscopy as a Lithographic Tool: Nanografting of DNA Nanostructures for Biosensing Applications . . . . . . . . . . . . . . . . . . . . . . Matteo Castronovo and Denis Scaini 16 Trapping and Immobilization of DNA Molecules Between Nanoelectrodes . . . . . Anton Kuzyk, J. Jussi Toppari, and Päivi Törmä 17 DNA Contour Length Measurements as a Tool for the Structural Analysis of DNA and Nucleoprotein Complexes . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Claudio Rivetti 18 DNA Molecular Handles for Single-Molecule Protein-Folding Studies by Optical Tweezers . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Ciro Cecconi, Elizabeth A. Shank, Susan Marqusee, and Carlos Bustamante 19 Optimal Practices for Surface-Tethered Single Molecule Total Internal Reflection Fluorescence Resonance Energy Transfer Analysis . . . . . . . . . . . . . . . . Matt V. Fagerburg and Sanford H. Leuba 20 Engineering Mononucleosomes for Single-Pair FRET Experiments . . . . . . . . . . . Wiepke J.A. Koopmans, Ruth Buning, and John van Noort 21 Measuring DNA–Protein Binding Affinity on a Single Molecule Using Optical Tweezers . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Micah J. McCauley and Mark C. Williams 22 Modeling Nanopores for Sequencing DNA . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Jeffrey R. Comer, David B. Wells, and Aleksei Aksimentiev

209 223

235

255

273 291

305 317

Index . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 359

Contributors Justin J. Abramson  •  Department of Mechanical Engineering, Columbia University, New York, NY, USA Aleksei Aksimentiev  •  Department of Physics, University of Illinois at Urbana-Champaign, Urbana, IL, USA Faisal A. Aldaye  •  Department of Systems Biology, Harvard Medical School, Boston, MA, USA Frank F. Bier  •  Department of Nanobiotechnology & Nanomedicine, Fraunhofer Institute for Biomedical Engineering, Branch Potsdam-Golm, Potsdam, Germany Ruth Buning  •  Leiden Institute of Physics, Leiden Universiteit, Leiden, The Netherlands Carlos Bustamante  •  Howard Hughes Medical Institute, Department of Physics, University of California, Berkeley, CA, USA Stephane Campidelli  •  CEA Saclay, Laboratoire d’Electronique Moléculaire, Gif-sur-Yvette Cedex, France Matteo Castronovo  •  Department of Biology, MONALISA Laboratory, College of Science and Technology, Temple University, PA, USA Ciro Cecconi  •  CNR-Istituto Nanoscienze S3, Department of Physics, University of Modena e Reggio Emilia, Modena, Italy Jennifer N. Cha  •  Department of Nanoengineering, UC San Diego, La Jolla, CA, USA Rahul Chhabra  •  University of Alberta, National Institute of Nanotechnology, Edmonton, AB, Canada Jeffrey R. Comer  •  Department of Physics, University of Illinois at Urbana-Champaign, Urbana, IL, USA Bernard A. Connolly  •  Chemical Nanoscience Laboratory, School of Chemistry, Newcastle University, Newcastle upon Tyne, UK Roberto Corradini  •  Dipartimento di Chimica Organica e Industriale, Univeristà di Parma, Parma, Italy Andrea Csaki  •  Institute of Photonic Technology (IPHT), Jena, Germany Matt V. Fagerburg  •  Departments of Cell Biology and Physiology and Bioengineering, University of Pittsburgh School of Medicine and Swanson School of Engineering, Petersen Institute of Nano Science and Engineering and University of Pittsburgh Cancer Institute, Pittsburgh, PA, USA Arianna Filoramo  •  CEA Saclay, Laboratoire d’Electronique Moléculaire, Gif-sur-Yvette Cedex, France Wolfgang Fritzsche  •  Institute of Photonic Technology (IPHT), Jena, Germany Alon A. Gorodetsky  •  Department of Chemistry, Columbia University, New York, NY, USA

ix

x

Contributors

Mark M. Green  •  Dipartimento di Chimica Organica e Industriale, Univeristã di Parma, Parma, Italy Graham D. Hamblin  •  Department of Chemistry, McGill University, Montreal, Canada Ralph Hölzel  •  Department of Nanobiotechnology & Nanomedicine, Fraunhofer Institute for Biomedical Engineering, Branch Potsdam-Golm, Potsdam, Germany James Hone  •  Department of Mechanical Engineering, Columbia University, New York, NY, USA Benjamin R. Horrocks  •  Chemical Nanoscience Laboratory, School of Chemistry, Newcastle University, Newcastle upon Tyne, UK Andrew Houlton  •  Chemical Nanoscience Laboratory, School of Chemistry, Newcastle University, Newcastle upon Tyne, UK Albert M. Hung  •  Department of Nanoengineering, UC San Diego, La Jolla, CA, USA Yonggang Ke  •  Dana-Farber Cancer Institute & Harvard Medical School, Boston, MA, USA Wiepke J.A. Koopmans  •  Leiden Institute of Physics, Leiden Universiteit, The Netherlands Alexander Kotlyar  •  Department of Biochemistry, The George S. Wise Faculty of Life Sciences, Tel Aviv University, Ramat Aviv, Israel Yamuna Krishnan  •  Biochemistry, Biophysics and Bioinformatics, National Centre for Biological Sciences, Bangalore, India Anton Kuzyk  •  Lehrstuhl für Bioelektronik, Physik-Department and ZNN/WSI, Technische Universität München, Garching, Germany Christian Leiterer  •  Institute of Photonic Technology (IPHT), Jena, Germany Sanford H. Leuba  •  Departments of Cell Biology and Physiology and Bioengineering, University of Pittsburgh School of Medicine and Swanson School of Engineering, Petersen Institute of NanoScience and Engineering, University of Pittsburgh Cancer Institute, Pittsburgh, PA, USA Chenxiang Lin  •  Dana-Farber Cancer Institute & Wyss Institute at Harvard University, Boston, MA, USA Yan Liu  •  Department of Chemistry and Biochemistry, The Biodesign Institute, Arizona State University, Tempe, AZ, USA Pik Kwan Lo  •  Department of Chemistry, McGill University, Montreal, Canada Rosangela Marchelli  •  Dipartimento di Chimica Organica e Industriale, Univeristà di Parma, Parma, Italy Susan Marqusee  •  Department of Molecular & Cell Biology, University of California, Berkeley, CA, USA Micah J. McCauley  •  Department of Physics, Northeastern University, Boston, MA, USA Christopher K. McLaughlin  •  Department of Chemistry, McGill University, Montreal, Canada Daisuke Miyoshi  •  Faculty of Frontiers of Innovative Research in Science and Technology (FIRST), and Frontier Institute for Biomolecular Engineering Research (FIBER), Konan University, Kobe, Japan

Contributors

Souvik Modi  •  Biochemistry, Biophysics and Bioinformatics, National Centre for Biological Sciences, Bangalore, India Khoa Nguyen  •  CEA Saclay, Laboratoire d’Electronique Moléculaire, Gif-sur-Yvette Cedex, France Colin Nuckolls  •  Department of Chemistry, Columbia University, New York, NY, USA Matteo Palma  •  Department of Mechanical Engineering & Applied Physics and Applied Mathematics, Columbia University, New York, NY, USA Andrew R. Pike  •  Chemical Nanoscience Laboratory, School of Chemistry, Newcastle University, Newcastle upon Tyne, UK Edda Reiß  •  Department of Nanobiotechnology & Nanomedicine, Fraunhofer Institute for Biomedical Engineering, Branch Potsdam-Golm, Potsdam, Germany Claudio Rivetti  •  Department of Biochemistry and Molecular Biology, University of Parma, Parma, Italy Denis Scaini  •  Sincrotrone Trieste, Basovizza, Trieste, Italy Stefano Sforza  •  Dipartimento di Chimica Organica e Industriale, Univeristà di Parma, Parma, Italy Elizabeth A. Shank  •  Harvard Medical School, Boston, MA, USA Jaswinder Sharma  •  Center for Integrated Nanotechnologies, Los Alamos National Laboratory, Los Alamos, NM, USA Michael P. Sheetz  •  Department of Biological Sciences, Columbia University, New York, NY, USA Friedrich C. Simmel  •  Physik Department, Technische Universität München, Munich, Germany Hanadi F. Sleiman  •  Department of Chemistry, McGill University, Montreal, Canada Thomas L. Sobey  •  Physik Department, Technische Universität München, Munich, Germany Naoki Sugimoto  •  Faculty of Frontiers of Innovative Research in Science and Technology (FIRST), and Frontier Institute for Biomolecular Engineering Research (FIBER), Konan University, Kobe, Japan Tullia Tedeschi  •  Dipartimento di Chimica Organica e Industriale, Università di Parma, Parma, Italy J. Jussi Toppari  •  Department of Physics, Nanoscience Center, University of Jyväskylä, Jyväskylä, Finland Päivi Törmä  •  Department of Applied Physics, School of science, Aalto University, Aalto, Finland John van Noort  •  Leiden Institute of Physics, Leiden Universiteit, Leiden, The Netherlands James Vesenka  •  Department of Chemistry and Physics, University of New England, Biddeford, ME, USA David B. Wells  •  Department of Physics, University of Illinois at Urbana-Champaign, Urbana, IL, USA

xi

xii

Contributors

Mark C. Williams  •  Department of Physics, Northeastern University, Boston, MA, USA Shalom J. Wind  •  Department of Applied Physics and Applied Mathematics, Columbia University, New York, NY, USA Hao Yan  •  Department of Chemistry and Biochemistry, The Biodesign Institute, Arizona State University, Tempe, AZ, USA Hua Yang  •  Department of Chemistry, University of British Columbia, Vancouver, Canada

Chapter 1 Synthesis and Characterization of Self-Assembled DNA Nanostructures Chenxiang Lin, Yonggang Ke, Rahul Chhabra, Jaswinder Sharma, Yan Liu, and Hao Yan Abstract The past decade witnessed the fast evolvement of structural DNA nanotechnology, which uses DNA as blueprint and building material to construct artificial nanostructures. Using branched DNA as the main building block (also known as a “tile”) and cohesive single-stranded DNA (ssDNA) ends to designate the pairing strategy for tile–tile recognition, one can rationally design and assemble complicated nanoarchitectures from specifically designed DNA oligonucleotides. Objects in both two- and three-dimensions with a large variety of geometries and topologies have been built from DNA with excellent yield; this development enables the construction of DNA-based nanodevices and DNA-template directed organization of other molecular species. The construction of such nanoscale objects constitutes the basis of DNA nanotechnology. This chapter describes the protocol for the preparation of ssDNA as starting material, the self-assembly of DNA nanostructures, and some of the most commonly used methods to characterize the self-assembled DNA nanostructures. Key words: DNA nanotechnology, Self-assembly, Electrophoresis, Atomic force microscopy

1. Introduction The notion that DNA is merely the gene encoder of living ­systems has been eclipsed by the successful development of DNA nanotechnology. DNA is an excellent nanoconstruction material because of its inherent merits: First, the rigorous Watson-Crick base-pairing makes the hybridization between DNA strands highly predictable. Second, the structure of the B-form DNA double helix is well-understood; its diameter and helical repeat have been determined to be ~2 and ~3.4 nm (i.e., ~10.5 bases), respectively, which facilitates the modeling of even the most complicated DNA nanostructures. Third, DNA possesses combined Giampaolo Zuccheri and Bruno Samorì (eds.), DNA Nanotechnology: Methods and Protocols, Methods in Molecular Biology, vol. 749, DOI 10.1007/978-1-61779-142-0_1, © Springer Science+Business Media, LLC 2011

1

2

Lin et al.

structural stiffness and flexibility. The rigid DNA double helixes can be linked by relatively flexible single-stranded DNA (ssDNA) to build stable motifs with desired geometry. Fourth, modern organic chemistry and molecular biology have created a rich toolbox to readily synthesize, modify, and replicate DNA molecules. Finally, DNA is a biocompatible material, making it suitable for the construction of multicomponent nanostructures made from hetero-biomaterials. The field of structural DNA nanotechnology began with Nadrian Seeman’s vision of combining branched DNA molecules bearing complementary sticky-ends to construct two-dimensional (2D) arrays (1) and his experimental construction of a DNA object topologically equal to a cube (2). Today, DNA self-assembly has matured with such vigor that it is currently possible to build micro- or even millimeter-sized nanoarrays with desired tile geometry and periodicity as well as any discrete 2D or 3D nanostructures we could imagine (3–8). Modified by functional groups, those DNA nanostructures can serve as scaffolds to control the positioning of other molecular species (9–21), which opens opportunities to study intermolecular synergies, such as protein–protein interactions, as well as to build artificial multicomponent nanomachines (22–24). Generally speaking, the creation of a novel DNA motif usually requires the following steps: (1) Structural modeling: physical and/or graphic models are used to help the design of a new DNA motif; (2) Sequence design: in this step, specific sequences are assigned to all ssDNA molecules in the model; (3) Experimental synthesis of the DNA nanostructure; and (4) Characterization of the DNA nanostructure. The first two steps are crucial to program the outcome of self-assembly and assisted by computer software (25–30). In this chapter, we are going to describe the experimental protocols involved in steps 3 and 4.

2. Material All chemicals are purchased from Sigma-Aldrich (St. Louis, MO) unless otherwise noted. All buffer solutions are filtered and stored at room temperature unless otherwise noted. 2.1. Denaturing Polyacrylamide Gel Electrophoresis for the Purification of Synthetic SingleStranded DNA

1. Synthetic ssDNA (Integrated DNA Techonologies, Coralville, IA) with designated sequences. 2. TBE buffer (1×): 89 mM Tris–boric acid, pH 8.0, 2 mM ethylenediaminetetraacetic acid disodium salt (EDTA-Na2). 3. 20% urea-acrylamide Mix: 20% acrylamide (19:1 acrylamide:bis, Bio-Rad Laboratories, Hercules, CA), 8.3 M urea in 1× TBE buffer.

Synthesis and Characterization of Self-Assembled DNA Nanostructures

3

4. 0% Urea-acrylamide Mix: 8.3 M Urea in 1× TBE buffer. 5. Ammonium persulfate (APS): prepare 10% water solution and store at 4ºC. 6. N,N,N,N¢-tetramethyl-ethylenediamine (TEMED, Bio-Rad). 7. Bromophenol blue (BB) or xylene cyanole FF (XC) (2×): prepare 0.1% w/v solution of the dye in 90% formamide solution containing 10 mM NaOH and 1 mM Na2EDTA. 8. Ethidium bromide: prepare 300 mL 0.1 mg/mL solution in a glass tray for gel staining. 9. Elution buffer (1×): 500  mM ammonium acetate, 10  mM magnesium acetate, 2 mM EDTA-Na2. 10. 1-Butanol and 100% Ethanol. 11. Spin X centrifuge tube filters (Corning, Lowell, MA). 2.2. Self-Assembly of DNA Nanostructures

1. Polyacrylamide Gel Electrophoresis (PAGE) purified ssDNA.

2.3. Non-denaturing PAGE for the Characterization of Self-Assembled DNA Nanostructures

1. Self-assembled DNA nanostructures.

2. TAE-Mg buffer (10×): 0.4  M Tris–acetic acid, pH 8.0, 125 mM magnesium acetate, 20 mM EDTA-Na2. 2. 40% acrylamide (19:1 acrylamide:bis, Bio-Rad Laboratories, Hercules, CA) solution. 3. Non-denaturing loading buffer (10×): 0.2% w/v bromophenol blue and xylene cyanole FF in 1× TAE-Mg buffer containing 50% v/v glycerol. 4. DNA ladder with suitable size (Invitrogen, Carlsbad, CA). 5. TAE-Mg buffer (1×), TEMED, and 10% APS solution (vide supra). 6. Stains-All: prepare 0.01% w/v Stains-All in 45% v/v formamide solution.

2.4. Atomic Force Microscope Imaging of Self-Assembled DNA Arrays

1. Self-assembled DNA nanostructures. 2. TAE-Mg buffer (1×) (vide supra). 3. Mica discs (Ted Pella, Inc) and Atomic Force Microscope (AFM) cantilevers of choice with integrated probes (such as NP-S from Veeco, Inc for imaging in liquids).

3. Methods 3.1. Denaturing PAGE Purification of Synthetic ssDNA

With advanced solid state synthesis chemistry, DNA synthesizer can generate DNA strands with designated sequences up to 200base long. However, a significant yield drop is normally associated with the synthesis of longer DNA strands. For example,

4

Lin et al.

if the yield for the addition of one nucleoside is 99%, the yield for the synthesis of a 100-mer ssDNA is only ~37%. Therefore, it is very important to purify the synthetic DNA strands that are longer than 30 bases to maximize the self-assembly yield in the next step. Effective ways to purify ssDNA less than 200-base long include high performance liquid chromatography (HPLC) and PAGE. Here, we discuss the protocol for denaturing PAGE purification of synthetic DNA strands. 1. Set up the gel assembly following the manufacturer’s instruction (we use a Hofer SE 600 Ruby from GE Healthcare) (see Note 1). 2. Mix proper volume of 20% and 0% Urea-acrylamide stock solution to prepare the acrylamide solution with desired concentration. Each gel needs ~35 mL acrylamide solution. For example, to make an 8% polyacrylamide gel, take 14 mL of 20% Urea-acrylamide stock and mix with 21 mL of 0% Ureaacrylamide stock. Stir thoroughly to mix well. For each gel, add 262 mL of 10% APS solution and 14.7 mL of TEMED. Stir thoroughly to mix well. 3. Quickly cast the gel using 35 mL pipette and insert the comb. Make sure no air bubble is trapped in the gel. Leave the gel at room temperature for at least 30 min to allow it solidifies. 4. Prepare the DNA sample. Add DI water to each dry samples to make 0.5 OD260/mL DNA solution. Take 4 OD of each sample (8 mL) into newly labeled tubes (see Note 2) and the rest of the samples should be stored at −20ºC. Add 2× denaturing dye to each sample (BB, XC, or both) and add water to adjust the final volume to 20 mL. Heat the sample at 90ºC for 5 min to denature the DNA strands (see Note 3). 5. When the gel has polymerized, remove the combs and attach the upper buffer chamber (UBC) to the gel assembly. Add 1 × TBE buffer (running buffer) to the UBC and rinse the wells thoroughly with glass pipette. Drain the UBC and add fresh running buffer to cover all the wells. 6. Load the samples to each well. Load 10 mL/well into the gel wells (generally 2 OD per lane) using the gel loading tips. Be careful not to flush the sample out of the well (see Note 4). 7. Carefully put the UBC and gel assembly into the lower buffer chamber (LBC) with ~3.5 L 1× TBE buffer. Add buffers into both UBC and LBC to the marked MAX lines (see Note 5). 8. Turn on the circulating water and set the temperature to 35ºC. Secure the lid of the gel box and connect the electrodes to a DC power supply. Make sure the polarity is correct. Run gel at constant current ~30–40 mA per gel for around 2–3 h depending on the length of the interested DNA fragments.

5

Synthesis and Characterization of Self-Assembled DNA Nanostructures

Table 1 The tracking dye migration on polyacrylamide denaturing gels (Dyes migrate to the same point as DNA strand of the indicated size in a denaturing polyacrylamide gel) Polyacrylamide concentration

5%

6%

8%

10%

12%

Bromophenol blue (bp)

35

26

19

12

8

Xylene cyanole FF (bp)

130

106

76

55

26

The tracking dye in the loading buffer provides a rough marker of the migration of DNA fragments (Table 1). 9. Turn off the power supply and circulating water. Lift the gel from the gel assembly and carefully put it into a glass try containing ~300 mL ethidium bromide (see Note 6). Stain the gel for 5 min and destain it for 5 min in distilled water. 10. In a dark room, lift the gel gently and put it on the UV transilluminator. Turn on the UV at wavelength of 302  nm, use razor blade to cut the major band out (see Note 7). Turn off the UV lamp, chop the band into small pieces, and collect the small gel blocks into Spin X centrifuge tube filters. Add 500 mL of elution buffer into each filter; shake in cold room (4ºC) overnight before proceed to the next step (see Note 8). 11. Centrifuge the Spin X tube filters (4,600 × g for 6  min) to separate the elution buffer from gel blocks. Add 1  mL of 1-butanol to the collected elution buffer, vortex the tube for 1 min, and centrifuge it at 600 × g for 1 min. After the spin, discard the upper layer of 1-butanol with pipette into waste bottle under venting hood. The 1-butanol washing extracts ethidium bromide and tracking dyes from the DNA sample. 12. Add in 1 mL ethanol to the DNA sample and mix well. Leave the mixture in −20ºC freezer for 30 min. Spin at 16,200 × g for 30 min at 4ºC to precipitate DNA. Pour out the ethanol and wash the DNA pellet with 70% v/v ice cold ethanol if desired. Centrifuge the tube at 16,200 × g for 10  min after ethanol washing and pour out all liquid. 13. Use a vacuum concentrator (we use a Vacufuge from Eppendorf, Westbury, NY) to dry the purified DNA sample for 1 h at 30ºC. Add in 50 mL distilled H2O, vortex for 1 min to dissolve the DNA sample. Measure the absorbance of the DNA solution at 260 nm (OD260) using a UV-Vis spectrometer (we employ a Biophotometer from Eppendorf) and convert the measured OD260 value to molar concentration using

6

Lin et al.

the extinction coefficient (e) of the DNA strand provided by the oligonucleotide vendor. Adjust the concentration of all purified DNA strand solution to 30 mM (or any other value to the experimenter’s convenience) by adding distilled H2O. Store all DNA samples in −20ºC freezer. 3.2. Anneal DNA Strands to SelfAssemble DNA Nanostructures

The formation of hydrogen bonded DNA complex is a selfassembly process. The DNA strands are mixed at stoichiometric molar ratio in a near-neutral buffer containing divalent cations (usually Mg2+), heated to denature and then gradually cooled to allow the ssDNA molecules to find their correct partners and adopt the most energy-favorable conformation. 1. Add stoichiometric amount of DNA strands into one 1.5 mL tube (or any other suitable tube size). Add 10× TAE-Mg buffer and distilled H2O to adjust the final concentration of each DNA strand to be 1 mM or any other desired concentration. Mix well and close the tube tightly. 2. This mixture is then heated on a heat block to 95ºC for 5 min and cooled to the desired temperature by the following protocol: 20 min at 65ºC, 20 min at 50ºC, 20 min at 37ºC, and if desired, 20 min at room temperature. 3. To assemble large DNA constructs, such as 2D arrays, slow annealing is desirable. In this case, the mixture is placed on a floating rack, transferred to a 2 L water bath, which is preheated to about 90°C and placed inside a Styrofoam box, and allowed to cool slowly to the desired temperature over the period of 2 days. This slow annealing process can also be carried out on a thermal cycler (see Note 9).

3.3. Non-denaturing PAGE for the Characterization of Self-Assembled DNA Nanostructures

Non-denaturing PAGE is an effective assay to characterize the self-assembled DNA supermolecules. Well-formed DNA nanostructure should migrate as a distinct band after electrophoresis. Non-denaturing PAGE also provides information regarding the yield of self-assembly. A typical gel image showing the correct formation of four helix DNA tile (31) is shown in Fig. 1. 1. Set up the gel assembly following the manufacturer’s instruction as described in step 1, Subheading 3.1 (we use a Hoefer SE 600 Ruby, GE Healthcare). 2. Prepare non-denaturing acrylamide mixture from 40% acrylamide (acrylamide:bis 19:1) stock, 10× TAE-Mg buffer and distilled H2O. The final mixture should contain 1× TAE-Mg buffer. For example, to make an 8% non-denaturing gel, mix 7 mL of 40% acrylamide stock, 3.5 mL of 10× TAE-Mg buffer, and 24.5 mL H2O. Stir thoroughly to mix well. For each gel, add 262 mL of 10% APS solution and 14.7 mL of TEMED. Stir thoroughly to mix well.

Synthesis and Characterization of Self-Assembled DNA Nanostructures

7

Fig. 1. Nondenaturing gel (8% polyacrylamide) of the 4-helix complex stained with Stains-All. Equimolar mixtures of 1 mM of each strand were annealed, and the electrophoresis was run at room temperature. Lane  M is a 100  bp DNA ladder. Lanes 1–8 contain complexes with partial combination of the component strands. Strands included in the annealing are indicated with a schematic drawing above the lane. Lane 9 corresponds to the full complex with all of the component strands.

3. Quickly cast the gel using 35 mL pipette and insert the comb. Make sure no air bubble is trapped in the gel. Leave the gel at room temperature for at least 2 h to allow it solidify (see Note 10). 4. When the gel has polymerized, remove the combs and attach the UBC to the gel assembly. Add 1× TAE-Mg buffer (running buffer) to the UBC and rinse the wells thoroughly with glass pipette. Drain the UBC and add fresh running buffer to cover all the wells. 5. Add 10× non-denaturing loading buffer to the preannealed DNA samples (finally, the DNA should be in 1× loading buffer). Vortex to mix well. Immediately load the DNA samples to each well using the gel loading tips. (Be careful not to flush the sample out of the well.) Take note about the sequence of the samples loaded. A DNA ladder with proper size should be loaded into a separate lane as a reference. 6. Immerse the gel assembly (together with UBC) to the 1× TAE-Mg buffer in the LBC. Add buffers into both UBC and LBC to the marked MAX lines. It is important not to disturb the sample when adding buffer to UBC. Add buffer gently along the side of the chamber. 7. Turn on the circulating water and set the temperature to 20ºC. Secure the lid of the gel box and connect the electrodes to a DC power supply. Make sure the polarity is correct. Run gel at constant voltage ~200 V for 4–8 h depending on the size of the interested DNA complexes.

8

Lin et al.

8. Turn off the power supply and circulating water. Lift the gel from the gel assembly and carefully put it into a glass tray containing ~300 mL 0.01% Stains-All solution. (Wear gloves all time when working with Stains-All). Low percentage gels can be fragile thus should be treated with extreme care. Stain the gel for 2 h and then rinse the gel well using distilled H2O. 9. Place a slide of transparency on the white lamp of the transluminator, carefully put the gel on it, and turn on white light to destain the gel until the gel appears almost colorless. This takes approximately 10–20 min. Watch the color change to avoid overdestain. 10. Use Kimwipe to wick the water off as much as possible, and then cover the gel with another transparency. Make sure no air bubble is trapped between the gel and the transparency. Also avoid stripes and pattern caused by thin layer of water. Scan the gel on a desktop scanner and save the image. 3.4. AFM Imaging of Self-Assembled DNA Arrays

1. The protocol described here assumes the use of PicoPlus AFM (Agilent). To start the imaging session, turn on the computer, Pico Scan controller, and then the AC controller. Open software “Pico Scan.” 2. Choose tapping mode AFM (AC AFM) and insert proper AFM tip into the tip holder on the top of the scanner. For AAC (acoustic AC) mode, use the gold-coated silicon nitride tip (NP-S tip, Veeco) for imaging in liquid or the proper acoustic AC tip (Veeco) for imaging in air. For the NP-S tips, use the tip on the thinner and shorter cantilever for imaging. 3. Sample preparation: Assemble a piece of freshly cleaved mica as the bottom of the fluid cell on the sample stage. Spot a 2 mL of 1 mM NiCl2 solution on mica and leave it to adsorb on the surface for 2 min. Then, add a 2 mL of the sample to the spot and leave it to adsorb on the surface for another 2 min. Finally, add 400 mL 1× TAE-Mg buffer onto the mica in the fluid cell. The Ni2+ adsorbed on mica surface can help the DNA array stay on the surface during the scanning. Attach the sample stage to the magnetic posts on the AFM. 4. Place the scanner on the sample stage with the tip pointing down. Lock the scanner. Turn on laser switch and plug in the detector. Move the laser spot so that it is on the back of the cantilever tip. Adjust the position of the photodiode inside the detector to maximize the sum of signal. Also make sure that the reflected laser spot is at the center of the photodiode. 5. Tune the tip and choose drive frequency with maximum amplitude. Set the parameters for scanning. Proportional gain and integral gain are 0.5 both or larger (94%) (30, 31). The submonomeric approach must be used to make sure that the C2-modified chiral PNA is synthesized with minimal racemization, which can be checked by chiral GC analysis (33). 4. For the insertion of chiral PNA monomers with stereogenic centers in position 5, which are not affected by racemization, and of achiral PNA monomers, normal SPPS procedures can be used (16b, 32). 5. Diethylthiadicarbocyanine solution: 500 mM 3,3′-diethylthiadicarbocyanine dye (Sigma–Aldrich St. Louis, MO, USA) in methanol. 6. Phosphate buffer for measurements: 10  mM phosphate ­buffer, pH 7.0.

3. Methods 3.1. Switching the DNA Handedness Through the B–Z Transition

The conditions able to induce a B–Z transition in a poly [d(CG)]2 and in a mixed sequence containing a d(CG)10 insert (see Note 2) in a B-DNA (Fig. 2a, d) domain are described (see Notes 3 and 4). 1. The concentration of DNA can be determined by spectrophotometry, according to the molar absorbance provided by the producer. For poly d(GC), the e257 of 8,400 mol−1 cm−1 must be used to calculate the concentration expressed as number of base residues independent of the molecular weight.

86

Corradini et al.

2. Dissolve DNA in a 80–100  mM concentration (in base) in cacodylate buffer (see Note 1), without the addition of salts. If no buffer is used, trace amounts of divalent cations must not be present (we used water with 18 MW × cm resistivity) (see Note 5). 3. Prepare a DNA solution without NaCl (or with 10  mM NaCl), and without any other additives (see Note 6). This solution is the reference B-DNA form (Fig. 2a, c). 4. Prepare a solution of poly d(CG) in the presence of 4–5 M NaCl (or 1 M MgCl2). Heat at 60°C for a few minutes and then slowly cool to room temperature. This will cause the formation of Z-DNA (Fig. 2b) (see Note 7). If a solution of poly d(me5CG) is used, 2  M NaCl is sufficient to obtain Z-DNA conversion (Fig. 2e). 5. Prepare a solution of mixed sequence containing a d(CG)10 insert in a B-DNA domain (as depicted in Fig.  1b) in the presence of 5  M NaCl, heat at 60°C for few minutes, and then slowly cool to room temperature. The resulting solution has the insert mostly in the Z-conformation, with the remaining part mostly in the B-conformation, with two B–Z junctions spanning from 3 to 10 bp each (Fig. 2c, d). 6. Record a UV spectrum of each solution between 200 and 350 nm at 25°C (or at the desired temperature). Slight differences are observed for the B- and Z-forms, with the latter having a lower absorption at 260 nm and presenting a shoulder in the 280–300-nm region. 7. Record a CD spectrum in the 200–350-nm range at 25°C. The B-form has a maximum at 280 nm, a crossover around 265–270 nm, and a minimum at 250 nm (plus other transitions more intense in the 200–230 range). The Z-form has a minimum in the 290–295-nm region, a crossover at 280 nm, and a maximum near 260–265 nm, plus other transitions more intense in the 200–230 range. In Fig. 2a, b, a typical B–Z transition is reported for d(CG)10 and d(CG)20, showing additivity of the base contribution. In Fig. 2 c, d, the transition obtained for a segment containing two B–Z junctions is reported. Assuming a typical spectrum of the B-DNA segments, the difference obtained by subtracting to the spectrum measured the contribution of the B-segments shows the typical profile and intensity of the Z-form of same length (see Fig. 2d). 8. B–Z transition. To a portion of the solutions in point 2, with 10  mM NaCl, add a solution of [Co(NH3)2]3+ to a final 20  mM concentration, under stirring (see Notes 8 and 9). This leads to a fast transition to the Z-form. The concentration of the [Co(NH3)2]3+ necessary to obtain a half transition

Control of Helical Handedness in DNA and PNA Nanostructures

87

is dependent on the concentration of NaCl (see Fig. 2f ), and increases as [Na+] increases (see Note 10). 9. Record changes in UV and CD spectra, in particular at 260 and 294 nm, and plot these as a function of salt concentration. A sigmoidal curve should be observed in each case, allowing for the detection of the mid-transition point, as illustrated by the graph reported in Fig. 2g. 3.2. Inducing Handedness in PNA Segments

1. Helix handedness prediction. Though most chiral singlestranded PNAs have low CD signals and, therefore, are not preorganized in helical structures, they show helical behavior when forming duplexes with the complementary achiral PNAs (Fig.  3c) (see Note 11). The CD results associated with helicity can be used to predict the preferred handedness of PNAs as a function of the stereochemistry of chiral monomers: (a) C2-substituted (synthesized from d-amino acid: 2D, Fig.  3a) or C5-substituted (synthesized from l-amino acid: 5L) or with both stereogenic centers (2D, 5L) simultaneously present: right-handedness preferred; (b) C2-substituted (from l-amino acid, 2L) or C5-substituted (from d-amino acid, 5D) or with both stereocenters (2L, 5D) simultaneously present: left-handedness preferred; (c) 2L and 5L stereocenters simultaneously present (2L, 5L): chiral conflict, but the induction exerted by the stereocenter in position 5 is stronger, thus right-handedness is preferred; (d) 2D and 5D stereocenters  (2D, 5D) simultaneously present: chiral conflict, but the  induction exerted by the stereocenter in position 5 is ­stronger, thus left-handedness is preferred. The preference for the ­helical sense caused by incorporation of a non-racemic chiral center at C2 and/or C5 will in turn affect the binding affinity of PNA for the nucleic acids (DNA or RNA) (see Note 12) (32). 2. Prepare a solution containing the chiral PNA (5 mM) and its complementary antiparallel achiral PNA strand (5  mM) in phosphate buffer for measurements (see Note 13). 3. Measure the circular dichroism spectrum between 220 and 350 nm. A prevalence of right-handed PNA helices is characterized by a negative Cotton effect at 275 nm and a higher (in absolute value) positive Cotton effect at 260 nm (full line in Fig. 4a). A prevalence of left-handed PNA helices is characterized by a positive Cotton effect at 275 nm and a higher (in absolute value) negative Cotton effect at 260 nm (dotted line in Fig. 4a). Though the exact percentages of right- and left-handed structures in PNA:PNA duplexes are not exactly known and are still under investigation, evidences indicate that at least for antiparallel PNA:PNA duplexes, two different PNA modifications can be compared as their ability to induce

88

Corradini et al.

a preferred handedness: if the length, the sequence, and the concentrations of the PNAs are the same, the intensity of the antiparallel CD spectra can be taken as a measure of the extent of the helical induction. 4. Prepare a solution containing the chiral PNA:PNA duplex (5 mM of each strand in phosphate buffer for measurements) in 10  mM phosphate buffer at pH 7. To this solution, add 3%  volume of diethylthiadicarbocyanine solution. The dye aggregates onto the PNA–PNA duplex and the handedness of the duplex is transferred to the dye aggregate (see Fig. 4c). 5. Measure the absorbance and circular dichroism spectrum between 400 and 700 nm. A shift in the absorption spectrum (from 648 to 534 nm) is observed if the PNA:PNA–dye interaction occurs. 6. A prevalence of right-handed PNA helices is characterized by a positive CD at about 540 nm and a negative one at about 520  nm (full line in Fig.  4b). A prevalence of left-handed PNA:PNA helices is characterized by opposite signs (dotted line in Fig. 4b) (see Note 14).

4. Notes 1. Other buffers that avoid the use of arsenic compounds are also suitable. We tested 5 mM Tris–HCl at pH 7.0, with similar results. 2. The occurrence of Z-DNA structures is strongly dependent on the sequence and base modification (34); the most favorable combinations for Z-DNA are repeated d(CG) sequences (35, 36). Unmodified d(CG)4, but not d(GC)4, undergoes the B–Z transition with a midpoint of transition at 3 M NaCl. For short sequences, the use of CG and not GC repeats is recommended. 3. The preference for the Z-forms varies as follows: d(CG) > d (TG)n > d(GGGC)n > d(TA)n (37). Substitution of cytosine with 5-methylcytosine (me5C) or other 5-substituted cytosines (bromo or iodo) has the effect of favoring the otherwise unstable Z-DNA structures, allowing this particular form to be obtained under conditions near normal physiological salt concentration. 4. In order to evaluate the possible effect of a molecule on the B–Z transition, select a condition under which B- and Z-forms are present in almost equal amounts. First, perform a study of the extent of the Z-form of the DNA as a function of salt concentration. Add the molecule under study to the solution

Control of Helical Handedness in DNA and PNA Nanostructures

89

in which B- and Z-forms are present in 1:1 ratio. Record changes in UV and CD spectra. If the 260-nm signal has increased and the 280-nm signal has decreased in UV, the extent of the Z-form has increased, and vice-versa for the B-form. The change in the percentage of B- and Z-forms can be better calculated by CD signals at 294 and 260 nm. 5. There are reports of Z-induction at very low salt concentration in the presence of traces of divalent cations. The effect of counterions has been thoroughly studied in the case of Ni2+induced B–Z transitions and the following order of Ni2+ concentration needed has been found: NaCl ≈ Me4NCl > LiCl >>  MgCl2 > no salt > NaBF4 ≈ NaNO3 ≈ NaClO4 (38). 6. Several additives can affect the B–Z transition depending on their chirality (15). Examples of Z-DNA inhibitors are actinomycin, adriamycin, and mitomycin; Z–B inducers are ethidium bromide, daunomycin, and netropsin. A more complete list is reported in Ref. (34). 7. The kinetics of formation of Z-DNA are slow. Therefore, incomplete conversion can lead to contrasting CD results. 8. Many cationic Z-DNA inducers at higher concentrations can cause DNA condensation (aggregation of several DNA segments), which competes with the formation of Z-DNA. Vigorous stirring is recommended. A UV spectrum from 350 to 200 nm should also be recorded in order to rule out the presence of a condensation process (in which case a drift of baseline is observed). 9. A solution of spermine added to a final 10 mM concentration can be used as an alternative to this method. Under stirring (see Note 8), it is heated at 60°C for 10 min and then slowly cooled to room temperature. This leads to conversion to the Z-form. 10. Other tools are available for detecting Z-DNA, such as gel electrophoresis, NMR, Z-DNA-specific antibodies (35), fluorescence energy transfer (FRET) (39), exciton chirality of appended porphyrins (40), and Raman spectroscopy (41). 11. The helical preference of a chiral PNA strand is most apparent when the PNA is in a well-defined helical conformation, i.e., when involved in a PNA–PNA double helix (with an achiral complementary PNA). Although in some cases the presence of a helical conformation might be hypothesized also in single-strand PNAs, caution is suggested when interpreting similarities in CD spectra or other experiments as “proofs” for a helical conformation of single-strand PNAs. 12. Usually, PNAs having a preference for left-handedness will nonetheless bind to DNA and RNA, by assuming the righthanded helical conformation dictated by the nucleic acids.

90

Corradini et al.

It  is exactly this forced unnatural conformation that is responsible for the decrease in stability in these PNA–DNA and PNA–RNA duplexes. 13. In order to avoid confusing effects, it is important that the complementary PNA is completely achiral, i.e., also amino acids linked at the C-terminus (as it is often the case with lysine) are to be avoided. 14. Aggregation of the dye onto the PNA–PNA duplexes depends on different factors, including steric hindrance and electrostatic repulsion or attraction, thus two different PNA modifications cannot be compared on their ability to induce a preferred handedness by considering the induced intensity of the dye in the CD spectra. References 1. (a) Hembury, G. A. , Borovkov, V.V. , and Inoue, Y. (2008) Chirality-Sensing Supramole­ cular Systems Chem Rev 108, 1–73. (b) Green, M. M. (2000) A model for how polymers amplify chirality. In Circular Dichroism (2nd Edition) Berova, N., Nakanishi, K., Woody, R.W. Eds Wiley-VCH, New York, 491–520. 2. (a) Green, M.M., Peterson, N. C., Sato, T., Teramoto, A., Cook, R., and Lifson, S. (1995) A helical polymer with a cooperative response to chiral information Science 268, 1860–6. (b) Green, M.M., Park, J.-W., Sato, T., Teramoto, A., Lifson, S., Selinger, R.L.B., and Selinger J.V. (1999). The macromolecular route to chiral amplification Angew Chem, Int Ed 38, 3139–54. 3. Tomar S., Green M. M., and Day L. A. (2007) DNA-Protein Interactions as the Source of Large Length Scale Chirality Evident in the Liquid Crystal Behavior of Filamentous Bacteriophages J Amer Chem Soc 129, 3367–75. 4. Hecht S.M. Ed. (1996) Bioorganic ChemistryNucleic Acids. Oxford University Press, Oxford-UK. 5. a) Seeman, N.C. (2007) An Overview of Structural DNA Nanotechnology Mol Biotechnol 37, 246–57. b) Brucale M., Zuccheri G., and Samorì B. (2006) Mastering the complexity of DNA nanostructures Trends in Biotech 24, 3427–34. 6. Mao, C.D., Sun, W.Q., Shen, Z.Y., and Seeman, N.C. (1999) A nanomechanical device based on the B-Z transition of DNA Nature 397, 144–6. 7. Du S. M., Stollar B. D., and Seeman, N.C. (1995) A Synthetic DNA Molecule in Three

8.

9. 10.

11.

12.

13.

14.

15.

Knotted Topologies J Am Chem Soc 117, 1194–1200. Michaud, M., Jourdan, E., Raavelet, C., Villet, A., Ravel, A., Grosset, C., and Peyrin, E. (2005) Immobilized DNA aptamers as target specific chiral stationary phases for resolution of nucleoside and amino acid derivative enantiomers Anal Chem 76, 1015–20. Roelfes, G., and Feringa, B.L. (2005) DNAbased asymmetric catalysis. Angew Chem Int Ed Engl 44, 3230–2. Li, X., and Liu, D. (2004) DNA-templated organic synthesis: nature’s strategy for controlling chemical reactivity applied to synthetic molecules Angew Chem Int Ed 43, 4848–70. Hannah, K.C., and Armitage, B.A. (2004) DNA-templated assembly of helical cyanine dye aggregates: a supramolecular chain polymerization Acc Chem. Res 37, 845–53. Shemer, G., Krichevski, O., Markovich, G., Molotsky, T., Lubitz, I., and Kotlyar, A.B. (2006) Chirality of Silver Nanoparticles Synthesized on DNA J Am Chem Soc 128, 11006–7. Dukovic, G., Balaz, M., Doak, P., Berova, N. D., Zheng, M., Mclean, R.S., and Brus, L.E. (2006) Racemic Single-Walled Carbon Nanotubes Exhibit Circular Dichroism When Wrapped with DNA J Am Chem Soc 128, 9004–5. Ha, S.C., Lowenhaupt, K., Rich A., Kim, Y.-G., and Kim, K.K. (2005) Crystal structure of a junction between B-DNA and Z-DNA reveals two extruded bases Nature 437, 1183–6. Corradini, R., Sforza, S., Tedeschi, T., and Marchelli R. (2007) Chirality as a Tool in

Control of Helical Handedness in DNA and PNA Nanostructures

16.

17.

18.

19.

20.

21.

22.

23.

24.

25.

26.

Nucleic Acid Recognition: Principles and Relevance in Biotechnology and in Medicinal Chemistry Chirality 19, 269–94. a) Nielsen, P.E., Egholm, M., Berg, R.H., and Buchardt, O. (1991) Sequence-Selective Recognition of DNA by Strand Displacement with A Thymine-Substituted Polyamide Science 254, 1497–1500. b) Nielsen P.E. (Ed.) (2004) Peptide Nucleic Acids: Protocols and Applications (Second Edition) Horizon Bioscience, Norfolk (UK). Corradini, R., Sforza, S., Tedeschi, T., Totsingan, F., and Marchelli, R. (2007) Peptide nucleic acids with a structurally biased backbone: effect of conformational constraints and stereochemistry Curr Top Med Chem 7, 681–94. Menchise, V., De Simone, G., Tedeschi, T., Corradini, R., Sforza, S., Marchelli, R., Capasso, D., Saviano, and M., Pedone, C. (2003) Insights into peptide nucleic acid (PNA) structural features: the crystal structure of a D-lysine-based chiral PNA–DNA duplex Proc Natl Acad Sci USA 100, 12021–6. Lukeman, P.S., Mittal, A.C., and Seeman, N.C. (2004) Two dimensional PNA/DNA arrays: estimating the helicity of unusual nucleic acid polymers Chem Commun 1694–5. Wittung, P., Eriksson, M., Lyng, R., Nielsen, and P. E., Norden, B. (1995) Induced Chirality in PNA-PNA Duplexes J Am Chem Soc 117, 10167–73. Totsingan, F., Jain, V., Bracken, W. C., Faccini, A., Tedeschi, T., Marchelli, R., Corradini, R., Kallenbach, N.R., and Green, M.M. (2010) Conformational Heterogeneity in PNA:PNA Duplexes Macromolecules 43, 2692–2703. Rasmussen, H., Liljefors, T., Petersson, B., Nielsen, P. E., and Kastrup, J. S. (2004) The influence of a chiral amino acid on the helical handedness of PNA in solution and in crystals J Biomol Struct Dyn 21, 495–502. Pino, P., and Luisi, P.L. (1968) Optical activity and conformation in stereoregular vinyl polymers J Chimie Physique Physico-Chimie Bio. 65, 130–9. Puschl, A., Sforza S., Haaima, G., Dahl, O., and Nielsen, P.E. (1998) Peptide nucleic acids (PNAs) with a functional backbone Tetrahedron Lett 39, 4707–10. Sforza, S., Haaima, G., Marchelli, R., and Nielsen, P.E. (1999) Chiral peptide nucleic acids (PNAs). Helical handedness and DNA recognition Eur J Org Chem 197–204. Sforza, S., Corradini, R., Ghirardi, S., Dossena, A., and Marchelli, R. (2000) DNA Binding

27.

28.

29.

30.

31.

32.

33.

34.

35.

36.

91

of  a D-Lysine-Based Chiral PNA: Direction Control and Mismatch Recognition Eur J Org Chem 2905–13. Smith, J.O., Olson, D.A., and Armitage B.A. (1999) Molecular Recognition of PNA-Containing Hybrids: Spontaneous Assembly of Helical Cyanine Dye Aggregates on PNA Templates J Am Chem Soc 121, 2686–95. Behe, M., and Felsenfeld, G. (1981) Effects of methylation on a synthetic polynucleotide: The B-Z transition in poly(dG-m5dC) poly(dG-m5dC) Proc Natl Acad Sci USA 78, 1619–23. Uhlmann, E., Peyman, A., Breipohl, G., and Will D.W. (1998) PNA: Synthetic Polyamide Nucleic Acids with Unusual Binding Properties Angew Chem Int Ed 37, 2796–2823. Sforza, S., Tedeschi, T., Corradini, R., Ciavardelli, D., Dossena, A., and Marchelli, R. (2003) Fast, Solid-Phase Synthesis of Chiral Peptide Nucleic Acids with a High Optical Purity by a Submonomeric Strategy Eur J Org Chem 1056–63. Tedeschi, T., Sforza, S., Maffei, F., Corradini, R., and Marchelli R. (2008) A Fmoc-based submonomeric strategy for the solid phase synthesis of optically pure chiral PNAs Tetrahedron Lett 49, 4958–61. Sforza, S., Tedeschi, T., Corradini, R., and Marchelli, R. (2007) Induction of Helical Handedness and DNA Binding Properties of  Peptide Nucleic Acids (PNAs) with Two Stereogenic Centres Eur J Org Chem 5879–85. Corradini, R., Di Silvestro, G., Sforza, S., Palla, G., Dossena, A., Nielsen, P.E., and Marchelli, R. (1999) Direct Enantiomeric Separation of N-aminoethyl amino acids: Determination of the Optical Purity of Chiral Peptide Nucleic Acids (PNAs) by GC Tetrahedron Asymm 10, 2063–6. Fuertes, M.A., Cepeda, V., Alonso, C., and Pérez, J.M. (2006) Molecular Mechanisms for the B − Z Transition in the Example of Poly[d(G − C)·d(G − C)] Polymers. A Critical Review Chem Rev 106, 2045–64. Pohl, F.M., and Jovin, T.M. (1972) Saltinduced co-operative conformational change of a synthetic DNA: equilibrium and kinetic studies with poly (dG-dC) J Mol Biol 67, 375–96. Wang, A.H., Quigley, G.J., Kolpak, F.J., Crawford, J.L., van Boom, J.H., van der Marel, G., and Rich, A. (1979) The molecular Structure of the Left-Handed Z-DNA Double Helix at 1.0 Angstrom Atomic Resolution. Nature 282, 680–6.

92

Corradini et al.

37. Herbert, A., and Rich, A. (1999) Left-handed Z-DNA: structure and function Genetica 106, 37–47. 38. Spingler B. (2005) Anions or Cations: Who Is in Charge of Inhibiting the Nickel(II) Promoted B- to Z-DNA Transition? Inorg Chem 44, 831–3. 39. Jares-Erijman, E.A., and Jovin, T.M. (1996) Determination of DNA Helical Handedness by Fluorescence Resonance Energy Transfer J Mol Biol 257, 597–617. 40. a) Balaz, M., De Napoli, M., Holmes, A.E., Mammana, A., Nakanishi, K., Berova, and N.,

Purrello, R. (2005) A Cationic Zinc Porphyrin as a Chiroptical Probe for Z-DNA Angew Chem Int Ed 44, 4006-9. b) Balaz, M., Li, B.C., Steinkruger, J.D., Ellestad, G.A., Nakanishi, K., and Berova, N. (2006) Porphyrins conjugated to DNA as CD reporters of the salt-induced B to Z-DNA transition Org Biomol Chem 4, 1865–7. 41. Dai, Z.Y., Thomas, G.A., Evertsz, E., and Peticolas, W.L. (1989) The length of a junction between the B and Z conformations in DNA is three base pairs or less Biochemistry 28, 6991–6.

Chapter 7 G-Quartet, G-Quadruplex, and G-Wire Regulated by Chemical Stimuli Daisuke Miyoshi and Naoki Sugimoto Abstract Guanine-rich DNA, which is widely distributed in the human genome, can fold into a supramolecular structure called the G-wire. The G-wire possesses promising characteristics as a functional element for various applications in vitro and in vivo. Here, we describe the preparative procedures for the G-wire and signatures of G-wire formation. Procedures for the regulation of G-wire formation by chemical stimuli will be useful for in vivo and in vitro applications. Key words:  G-wire, G-quadruplex, G-quartet, Guanine, Molecular crowding, Metal ion, Polymorphism

1. Introduction There are 3.2 billion base pairs in the entire human genome, most of which may form the canonical B-form duplex via sequence specific Watson–Crick base pairs (1). Although the biological function of the genome is the storage of genetic information, sequence-specific formation of B-form duplexes is also useful for the construction of nanostructures (2, 3). In fact, many researchers have demonstrated that sequence-specific duplexes can be used to create one-dimensional, two-dimensional, and three-dimensional nanostructures (4–8). To facilitate the practical use of DNA ­nanostructures, functionalization of the nanostructures is an important topic in DNA nanotechnology. The Human Genome Project revealed that a very small fraction of the genome contains protein-encoding sequences (1). Our understanding of the functions of the remaining noncoding

Giampaolo Zuccheri and Bruno Samorì (eds.), DNA Nanotechnology: Methods and Protocols, Methods in Molecular Biology, vol. 749, DOI 10.1007/978-1-61779-142-0_7, © Springer Science+Business Media, LLC 2011

93

94

Miyoshi and Sugimoto

f­raction is rapidly increasing. It has also been revealed that ­repetitive DNA sequences widely distributed in the human genome have the potential to fold into noncanonical structures. These noncanonical DNA structures can undergo structural transitions depending on their sequences or surrounding conditions, which in turn can control gene expression (9–11). The polymorphic nature of noncanonical DNA structures is useful for designing functional DNA nanostructures. The most common and frequently observed repetitive DNA sequences throughout the genomes of most organisms are Guanine rich (G-rich) sequences and their complementary Cytosine-rich (C-rich) strands (12, 13). Especially, it is well known that G- and C-rich sequences are found at the ends of the chromosomes (telomeres). G- and C-rich telomeric sequences can fold into G-quadruplexes and i-motifs, respectively. G-quadruplexes can be formed by intermolecular or intramolecular association with four Hoogsteen-paired coplanar guanines called a G-quartet (Fig. 1a) (14). The G-quadruplex, which can be formed by intermolecular or intramolecular association of G-rich strands in antiparallel or ­parallel orientations (Fig.  1b), shows high structural polymorphism depending on its surrounding conditions. For example, Sugimoto and coworkers reported that only 1  mM of divalent cations largely  affects the thermodynamic parameters of the

a

b H

R

N

N

G

H

N H N

G

R N H

H

G

O

N H

N

N

N H

O H

N R

N

H

O N

H

M+

H

N

N

N

O

H

H

N

G N

N H H

N H

N R

Fig. 1. (a) Chemical structure of G-quartet. Center circle indicates metal ion. (b) Structure of antiparallel (top) and parallel (bottom) G-quadruplexes.

G-Quartet, G-Quadruplex, and G-Wire Regulated by Chemical Stimuli

95

­antiparallel G-quadruplex of d(G4T4G4) (15). G-quadruplexes are more polymorphic under molecular crowding conditions. Molecular crowding causes d(G4T4G4) and d((G4T4)3G4) to undergo transitions from antiparallel to parallel G-quadruplexes (Fig.  2a) (16). Moreover, it was demonstrated that a duplex formed of G- and C-rich DNAs under uncrowded conditions

Fig. 2. (a) Structural transition of d(G4T4G4) and d(G4T4)3G4 induced by molecular ­crowding. (b) Structure of G-rich and C-rich strands under dilute and molecular crowding conditions. (c) Schematic illustration for G-wire formation.

96

Miyoshi and Sugimoto

­ issociates upon molecular crowding and that the G- and C-rich d DNAs individually fold into quadruplexes (Fig.  2b) (17). Importantly, it was shown that the ­parallel G-quadruplex induced by molecular crowding was a high-order DNA nanostructure based on G-quartets called a G-wire (16). The G-wire consists of numerous strands of guanine-rich DNA strands aligned in parallel (Fig.  2c). Guanine monomers including guanosine and its synthetic derivatives, and guanine ­oligomers from several to thousands of consecutive guanines have been utilized for G-wire formations (17). Chen reported that Sr2+ can induce the formation of a G-wire by d((G4T2)3G4) (18). Using atomic force microscopy (AFM), Henderson and coworkers directly observed for the first time that d(G4T2G4) forms a G-wire in the presence of Mg2+ and spermidine (19). They also found that the G-wire is a one-dimensional nanowire with a uniform height and width, and few bends or kinks (20). Kotlyar et  al. developed a long G-wire using a polyG (~3,000 bases) strand that is prepared enzymatically with the Klenow fragment (21, 22). Macgregor and coworkers used d(NxGy) or d(GyNx), where y > 10 and x > 5, to prepare a DNA nanowire that they called a frayed wire (23, 24). Sugimoto and coworkers reported that Ca2+ induces G-wire formation of d(G4T4G4) (25). They further demonstrated that a single G to A substitution in the loops leads to a drastically different structure. Under molecular crowding conditions, which is one of the most drastic differences between chemical conditions of a ­living cell and test tube, d(T2(G3T2G)3G) folds into very long G-wires, whereas d((G3T2A)3G3) folds into an antiparallel G-quadruplex (26). Notably, studies on G-wires further showed that they have useful characteristics for various applications. For example, there is growing interest in G-rich sequences as functional ­elements in molecular electronics (27), since theoretical study suggests that the G-wire structure is promising for nanoscale biomolecular electronics (28). For such electronic applications, it is important that the G-wire tends to deposit on a surface with orientational distribution with three preferential directions (29). It is well known that the G-wire has promise for effecting separations and transportations of cations (30). The characteristics of G-wires are promising not only for material applications but also for biological applications. For example, it was demonstrated that cellular uptake and nuclear localization of the G-wire are more efficient than that of a single-stranded DNA, allowing us to utilize a G-wire for a drug delivery system (31). Moreover, G-wire assembly into a liquid crystalline phase (32) was applied for NMR study of detergent-solubilized membrane proteins (33).

G-Quartet, G-Quadruplex, and G-Wire Regulated by Chemical Stimuli

97

2. Materials 2.1. Preparation of DNA Samples

1. The phosphoramidite monomers, solid support (controlled pore glass bead), and all reagents and solvents for DNA ­oligonucleotide synthesis from Glen research (Sterling, VA). 2. Poly-Pak (Glen research) and Sep-Pak (Waters) C18 reversedphase cartridges for deprotection and crude preparation of oligonucleotides. 3. Reagents for purification by reverse-phase high performance liquid chromatography (HPLC) from Wako Pure Chemical Industries (Osaka, Japan). Reverse-phase column for HPLC from Tosoh (Tokyo, Japan). 4. G-wire buffer 1: 100 mM NaCl, 50 mM MES (pH 6.1), and 40 wt% polyethylene glycol (PEG 200 with average molecular weight 200) or PEG 2000. 5. G-wire buffer 2: 50 mM MES (pH 6.1), 100 mM NaCl, and 20 mM CaCl2.

2.2. Polyacrylamide Gel Electrophoresis

1. 10× Tris–borate–EDTA (TBE) buffer (1  L): 890  mM Tris, 890 mM boric acid, 20 mM Na2EDTA, and ddH2O to adjust the volume to 1 L. Stored at room temperature. 2. 20% Acrylamide solution (1 L): 190 g acrylamide, 10 g bisacrylamide, 100 mL 10× TBE buffer, and ddH2O to adjust the volume to 1 L. Stored at 4°C (see Note 1). 3. 10% Ammonium persulfate (APS) (1 mL): 100 mg APS and ddH2O to adjust the volume to 1 mL. Stored at 4°C. 4. Gel loading buffer dye for non-denaturing PAGE (5  mL): 40 wt% glycerol, 0.2 wt% Blue dextran, 0.5 mL 10× TBE buffer, and ddH2O to adjust the volume to 5 mL.

2.3. UV/Vis and Circular Dichroism Measurement

1. UV measurement: Appropriate buffer, quartz cuvettes with 1.0- and 0.1 cm path lengths, and temperature controller.

2.4. Atomic Force Microscope Measurement

1. Mica: K2O·Al2O3·SiO2 from Ted Pella Inc. (Redding, CA).

2. Circular dichroism (CD) measurement: Appropriate buffer, quartz cuvettes with 1.0- and 0.1 cm path lengths, and temperature controller.

2. AFM probe: RTESP for tapping mode in air and NP-S for tapping mode in liquid, from Veeco Instruments (Plainview, NY). 3. AFM specimen buffer: 100  mM NaCl, 50  mM MES (pH 6.1), and 100 mM CaCl2.

98

Miyoshi and Sugimoto

3. Methods 3.1. Preparation of DNA Samples

1. DNA is synthesized on solid supports using the standard ß-cyanoethyl phosphoramidite methods (34). 2. The synthesized DNA oligonucleotides containing the 5¢-end dimethoxytrityl (DMT) groups are removed from the solid support, and base blocking groups are removed by treatment with concentrated 25% ammonia at 55°C for 8 h. After drying in vacuum, the oligonucleotides are passed through a Poly-Pak cartridge with 2% trifluoroacetic acid to remove the 5¢-end DMT groups. After deblocking operations, the oligonucleotides are desalted through a C-18 Sep-Pak cartridge column. 3. The oligonucleotides are purified by HPLC on a TSK-gel Oligo DNA RP column (Tosoh) with a linear gradient of 0–50% MeOH/H2O containing triethylammonium acetate (pH 7.0) (see Note 2). The final purities of the oligonucleotides are confirmed to be >98% by HPLC. The purified oligonucleotides are desalted again with a C-18 Sep-Pak cartridge before use. 4. DNA synthesis is confirmed by matrix-assisted laser desorption/ionization time-of-flight (MALDI-TOF) mass spectrometry (see Note 3). 5. Single-strand concentrations of the DNA oligonucleotides are determined by measuring the absorbance at 260 nm at a high temperature using a Shimadzu 1700 spectrophotometer (Shimadzu, Kyoto, Japan) connected to a thermoprogrammer. Single-strand extinction coefficients are calculated from mono­ nucleotide and dinucleotide data using the nearest-neighbor approximation (35). 6. To induce G-wire formation of d(G4T4G4) and d(G4T4)3G4, appropriate buffers are G-wire buffers 1 and 2 (see Note 4). 7. Before the measurement, the sample is heated to high temperature (typically around 80–95°C), gently cooled at a rate of 0.1–3.0°C/min, and incubated at the desired temperature for several hours (see Note 5).

3.2. UV Analysis

1. The UV absorbance is measured with a Shimadzu 1700 spectrophotometer (Shimadzu, Kyoto, Japan) equipped with a temperature controller. 2. Melting curves of G-wire and G-quadruplex are obtained by measuring the UV absorbance at 295 nm in appropriate buffers (see Note 6). 3. Confirm that the heating rate does not affect the shape of the melting curve. Generally, the heating rate for G-wire and

G-Quartet, G-Quadruplex, and G-Wire Regulated by Chemical Stimuli

99

Fig. 3. (a) Normalized UV melting curves for a typical G-quadruplex in a buffer of 100 mM KCl, 10 mM K2HPO4 (pH 7.0), and 1 mM K2EDTA containing 0, 10, 20, 30, or 40 wt% PEG 200 (from left to right ). (b) Differential UV spectrum of a typical G-quadruplex.

G-quadruplex is 0.1–0.5°C/min or possibly slower. DNA concentration is adjusted to an absorbance of around 1.0 at 260 nm. 4. The melting temperature (Tm) and thermodynamic parameters (enthalpy change, ∆H ˚; entropy change, T∆S ˚; and free energy change, ∆G ˚) for the G-wire and G-quadruplex formation are obtained from the UV melting curve (see Note 7). 5. The hypochromicity in the UV melting curve at 295 nm is observed for G-wire and G-quadruplex structures (Fig.  3a) (36). On the contrary, the duplex shows hyperchromicity at 295 nm. 6. The differential UV spectrum (UV spectra at a temperature higher than Tm − UV spectra at a temperature lower than Tm) shows a negative peak at around 295 nm in the case of G-wire and G-quadruplex structures (Fig. 3b) (37). 3.3. CD Analysis

1. CD experiments are carried out with a JASCO J-820 spectropolarimeter (JASCO, Hachioji, Japan). The temperature of the cell holder is regulated by a JASCO PTC-348 temperature controller (see Note 6). 2. The CD spectra are obtained by taking the average of at least three scans made from 200 to 350 nm. The buffer spectrum is subtracted from the DNA spectrum. 3. The CD spectra of the G-wire and parallel G-quadruplex have positive and negative peaks at around 260 and 240  nm, respectively (Fig. 4a). On the contrary, the CD spectra of the antiparallel G-quadruplex have positive and negative peaks at

100

Miyoshi and Sugimoto

Fig.  4. (a) CD spectra of 50  mM d(G4T4G4) in buffers containing 100  mM NaCl, 10  mM Na2HPO4 (pH 7.0), and 1  mM Na2EDTA without (bottom) or with 40 wt% co-solute at 4°C. Co-solutes used here are PEG 2000; PEG 200; tri(ethylene glycol), di(ethylene glycol), or ethylene glycol; and diEG (from top to second bottom at 260 nm). (b) Non-denaturing PAGE of d(G4T4G4) in the presence of 0, 10, 20, 30, or 40 wt% PEG 2000. Lane 1: 10-bp DNA marker, lane 2: single-stranded 24 mer DNA, lane 3: d(G4T4G4) with Mg2+ and spermidine, lanes 4–8: d(G4T4G4) with 0, 10, 20, 30, or 40 wt% PEG 2000.

around 295 and 260  nm. In addition, the mixed parallel G-quadruplex, in which three strands are in a parallel and one in an antiparallel direction, shows positive peaks at around 295 and 260 nm. 3.4. PAGE Analysis

1. The following instruction is for the Atto AE6220 gel system. This is easily adaptable to other systems. It is important to use clean glass plates. The glass plates are scrubbed with a rinsable detergent after use, rinsed extensively with distilled water, and dried. 2. Prepare a non-denaturing gel with 15 cm × 15 cm glass plates and 1.0  mm-thick spacer. For a non-denaturing acrylamide gel of 15  cm × 15  cm × 1.0  mm, 20  ml of gel solution is ­sufficient. For a 20% acrylamide gel, quickly mix 20  mL ­acrylamide solution, 180  mL APS, and 18  mL N,N,N ¢,N ¢tetramethylethylenediamine (TEMED); pour into the space between the two glass plates; insert the comb around 1 cm high; and leave it for 30–60 min (see Note 8). 3. After polymerization is complete, remove the comb and bottom spacers. Wash the gel plates with distilled water to remove any spilled acrylamide gel. 4. Fill the lower reservoir of the electrophoresis system with 1× TBE buffer. Set the gel plates to the gel system and fill the reservoir of the electrophoresis system with 1× TBE buffer until all of the sample wells are covered (see Note 9).

G-Quartet, G-Quadruplex, and G-Wire Regulated by Chemical Stimuli

101

5. Wash the wells with the filled 1× TBE buffer. Mix the loading buffer (5 mL) with 5 mL of sample and load 10 mL of each sample in a well. 6. Complete assembly of the gel system and connect to a power supply. The gel can be run for several ten minutes to several hours at 5 V/cm. 7. In order to stop running, the gel system is disconnected from the power supply and disassembled. The stacking gel with glass plates is decartelized and the gel is removed from the glass plate. 8. The separating gel is then immersed for 10 min in a dye solution such as GelStar nucleic acid gel stain (Cambrex, ME), SYBR gold (Invitrogen, Eugene, OR), or Stains-All (Sigma– Aldrich Co, St. Louis, MO) (see Note 10). 9. Pick up and immerse the gel in 1 × TBE buffer for 10 min to remove dyes binding to the gel. 10. In order to quantify the electrophoresis result, fluorescence intensity is analyzed with an image analyzer such as a Fujifilm FLA-5001 (Fujifilm, Tokyo, Japan). 11. Migration of the G-quadruplex is faster than that of a duplex of the same length. On the contrary, smearing and slower migration, including a ladder pattern, are observed for the G-wire (Fig. 4b). In addition, it is sometimes impossible for a larger G-wire to migrate through the gel (see Note 11). 3.5. AFM Measurement

1. A 25 mM DNA sample is dissolved in AFM specimen buffer and heated to 90°C for 10  min, gently cooled at a rate of 1.0°C/min, and incubated at 45°C for 3 h (see Note 12). 2. A 1 mL sample is deposited onto freshly cleaved mica three times, washed with 500 mL deionized water three times, and dried with a stream of N2 gas (see Note 13). 3. AFM images are obtained in the tapping mode using a Nanoscope III (Digital Instruments Inc., USA). The AFM image shows that the G-wire is a linear polymer, and the height and width of the G-wires are uniform with a few bends or kinks. Clear AFM images for G-wires were reported (38, 39).

4. Notes 1. Wear safety glasses, mask, and gloves since unpolymerized acrylamide is neurotoxic. 2. HPLC-grade water and methanol are recommended.

102

Miyoshi and Sugimoto

3. The matrix routinely used in MALDI-TOF mass spectrometry to analyze oligomer DNA is 3-hydroxypicolinic acid (3-HPA). 4. The pH value of buffers is adjusted with HCl and NaOH, KOH, or LiOH depending on the monovalent cation used. 5. Note that the highest temperature should be lower than the boiling temperature of the co-solute in the buffer, but higher than the melting temperature of the DNA structure. 6. When the measurement is carried out below room temperature, the cuvette-holding chamber is flushed with a constant stream of dry N2 gas to avoid condensation of water on the cuvette exterior. 7. A two-state transition between an unstructured random coil and structured states is required to evaluate the thermodynamic parameters. In the case of the G-wire, it is generally difficult to assume a two-state transition since the G-wire is not a homogeneous structure. 8. Remove air bubbles from the acrylamide gel solution before polymerization. Since the polymerization is an exothermic reaction, the completion of the reaction is detectable by ­temperature change. 9. Place the gel into the lower tank at an angle to avoid air ­bubbles forming at the bottom of the gel plates. 10. Wear safety glasses, mask, and gloves since these dyes are toxic. 11. Since the G-wire is thermally very stable, slower migration is often observed even in a denaturing gel. 12. Divalent metal ions such as Mg2+, Ca2+, or Ni2+ are required for immobilization of the DNA sample onto the mica surface, since freshly cleaved mica has a negative charge. Thus, TAE/Mg buffer (10× TAE–Mg (Tris–acetate–Mg2+) is 125  mM Mg–acetate, 400 mM Tris–HCl, and 10 mM Na2EDTA) can alternatively be used for AFM measurements of DNA nanostructure. 13. The preparation procedure for AFM measurements in liquid is different from that in air.

Acknowledgments This work was supported in part by Grants-in-Aid for Scientific Research, the Academic Frontier Project (2004–2009), and the Core Research project (2009–2014) of the Ministry of Education, Culture, Sport, Science and Technology (MEXT) of Japan; by the Long-range Research Initiative; and by the Hirao Taro Foundation of the Konan University Association for Academic Research.

G-Quartet, G-Quadruplex, and G-Wire Regulated by Chemical Stimuli

103

References 1. International Human Genome Sequencing Consortium (2004) Finishing the euchromatic sequence of the human genome. Nature 431, 931–945. 2. Seeman, N. C. (2003) DNA in a material world. Nature 421, 427–431. 3. Jaeger, L. and Chworos, A. (2006) The architectonics of programmable RNA and DNA nanostructures. Curr. Opin. Struct. Biol. 16, 531–543. 4. Sharma, J., Chhabra, R., Cheng, A., Brownell, J., Liu, Y., and Yan, H. (2009) Control of self-assembly of DNA tubules through ­integration of gold nanoparticles. Science 323, 112–116. 5. Liu, Y., Kuzuya, A., Sha, R., Guillaume, J., Wang, R., Canary, J. W., and Seeman, N. C. (2008) Coupling across a DNA helical turn yields a hybrid DNA/organic catenane doubly tailed with functional termini. J. Am. Chem. Soc. 130, 10882–10883. 6. He, Y., Ye, T., Su, M., Zhang, C., Ribbe, A. E., Jiang, W., Mao, C. (2008) Nature 452, 198–201. 7. Rothemund, P. W. (2006) Folding DNA to create nanoscale shapes and patterns. Nature 440, 297–302. 8. Goodman, R. P., Schaap, I. A. T., Tardin, C. F., Erben, C. M., Berry, R. M., Schmidt, C. F., and Turberfield, A. J. (2005) Rapid chiral assembly of rigid DNA building blocks for molecular nanofabrication. Science 310, 1661–1665. 9. Kumari, S., Bugaut, A., Huppert, J., and Balasubramanian, S. (2007) An RNA G-quadruplex in the 5’ UTR of the NRAS proto-oncogene modulates translation. Nat. Chem. Biol. 3, 218–221. 10. Rankin, S., Reszka, A. P., Huppert, J., Zloh, M., Parkinson, G. N., Todd, A. K., Ladame, S., Balasubramanian, S., and Neidle, S. (2005) Putative DNA quadruplex formation within the human c-kit oncogene. J. Am. Chem. Soc. 127, 10584–10589. 11. Qin. Y and Hurley, L. H. (2008) Structures, folding patterns, and functions of intramolecular DNA G-quadruplexes found in eukaryotic promoter regions. Biochimie, 90, 1149–1171. 12. Scaria, V., Hariharan, M., Arora, A., and Maiti, S. (2006) Quadfinder: server for identification and analysis of quadruplex-forming motifs in nucleotide sequences. Nucleic Acids Res. 34 (Web Server issue), W683–685.

13. Du, Z., Zhao, Y., and Li, N. (2008) Genomewide analysis reveals regulatory role of G4 DNA in gene transcription Genome Res. 18, 233–241. 14. Gellert, M., Lipsett, M. N., and Davies, D. R. (1962) Helix formation by guanylic acid. Proc. Natl. Acad. Sci. USA, 48, 2013–2018. 15. Miyoshi, D., Nakao, A., Toda, T., and Sugimoto, N. (2001) Effect of divalent cations on antiparallel G-quartet structure of d(G4T4G4). FEBS Lett. 496, 128–133. 16. Miyoshi, D., Nakao, A., and Sugimoto, N. (2002) Molecular crowding regulates the structural switch of the DNA G-quadruplex. Biochemistry 41, 15017–15024. 17. Miyoshi D, Matsumura S, Nakano S, Sugimoto N. (2004) Duplex dissociation of telomere DNAs induced by molecular crowding. J. Am. Chem. Soc. 126, 165–169. 18. Davis, J. T. (2004) G-quartets 40 years later: from 5’-GMP to molecular biology and supramolecular chemistry. Angew. Chem. Int. Ed. 43, 668–698. 19. Chen, F. M. (1992) Sr2+ facilitates intermolecular G-quadruplex formation of telomeric sequences. Biochemistry, 21, 3769–3776. 20. Marsh, T. C. and Henderson, E. (1994) G-wires: self-assembly of a telomeric ­oligonucleotide, d(GGGGTTGGGG), into large superstructures. Biochemistry 33, 10718–10724. 21. Marsh, T. C., Vesenka, J., and Henderson, E. (1995) new DNA nanostructure, the G-wire, imaged by scanning probe microscopy. Nucleic Acids Res. 23, 696–700. 22. Kotlyar, A., Borovok, N., Molotsky, T., Cohen, H., Shapir E., and Porath, D. (2005) Long Monomolecular G4-DNA Nanowires”. Adv. Mat. 17, 1901–1905. 23. Borovok, N., Molotsky, T., Ghabboun, J., Porath, D., and Kotlyar, A. (2008) Efficient procedure of preparation and properties of long uniform G4-DNA nanowires. Anal. Biochem. 374, 71–78. 24. Protozanova, E. and Macgregor, R. B. Jr. (1996) Frayed wires: a thermally stable form of DNA with two distinct structural domains. Biochemistry 35, 16638–16645. 25. Yanze, M. F., Lee, W. S., Poon, K., PiquetteMiller, M., and Macgregor, R. B. Jr. (2003) Cellular uptake and metabolism of DNA frayed wires. Biochemistry 42, 11427–11433. 26. Miyoshi, D., Nakao, A., and Sugimoto, N. (2003) Structural transition from antiparallel

104

27.

28.

29. 30.

31.

32. 33.

Miyoshi and Sugimoto to parallel G-quadruplex of d(G4T4G4) induced by Ca2+. Nucleic Acids Res. 31, 1156–1163. Miyoshi, D., Karimata, H., and Sugimoto, N. (2005) Drastic effect of a single base ­difference between human and tetrahymena telomere sequences on their structures under molecular crowding conditions. Angew. Chem. Int. Ed. 44, 3740–3744. Porath, D., Bezryadin, A., de Vries, S., and Dekker, C. (2000) Direct measurement of electrical transport through DNA molecules. Nature 403, 635–638. Calzolari, A., Felice, R. D., Molinari, E. (2002) G-quartet biomolecular nanowires. Appl. Phys. Lett. 80, 3331–3333. Vesenka, J., Bagg, D., Wolff, A., Reichert, A., Moeller, R., and Fritzsche, W. (2007) Autoorientation of G-wire DNA on mica. Colloids Surf B Biointerfaces 58, 256–263. Davis, J. T. and Spada, G. P. (2007) Supramolecular architectures generated by self-assembly of guanosine derivatives. Chem. Soc. Rev. 36, 296–313. Spada, G. P. and Gottarelli, G. (2004) Synlett 596–602. Lorieau. J., Yao, L., and Bax, A. (2008) Liquid crystalline phase of G-tetrad DNA for NMR

34. 35.

36. 37.

38.

39.

study of detergent-solubilized proteins. J. Am. Chem. Soc. 130, 7536–7537. Sanger. W. (1984) Principle of Nucleic Acids and Structure, Springer-Verlag, New York. Richards, E. G. (1975) Use of tables in calculation of absorption, optical rotatory dispersion and circular dichroism of polyribon ucleotides. In Fasman,G.D. (ed.), Handbook of Biochemistry and Molecular Biology, 3rd edn. CRC Press, Cleveland, OH, USA, Vol. 1, pp. 596–603. Mergny, J. L., Phan, A. T., and Lacroix L. (1998) Following G-quartet formation by UV-spectroscopy. FEBS Lett. 435, 74–78. Miyoshi, D., Nakamura, K. Karimata, H., Ohmichi, T., and Sugimoto, N. Hydration of Watson-Crick base pairs and dehydration of Hoogsteen base pairs inducing structural polymorphism under molecular crowding conditions. J. Am. Chem. Soc., 130, in press (2009). Mergny, J. L., Li, J., Lacroix, L., Amrane, S., and Chaires, J. B. (2005) Thermal difference spectra: a specific signature for nucleic acid structures. Nucleic Acids Res. 33, e138. Kunstelj, K., Federiconi, F., Spindler, L., and Drevensek-Olenik, I. (2007) Self-organization of guanosine 5’-monophosphate on mica. Colloids Surf B Biointerfaces 59, 120–127.

Chapter 8 Preparation and Atomic Force Microscopy of Quadruplex DNA James Vesenka Abstract The purpose of this chapter is to provide detailed instructions for the preparation and atomic force microscopy (AFM) imaging of linear chains of quadruplex DNA (a.k.a. “G-wire DNA”). Successful selfassembly of long chain quadruplex DNA requires pure concentrated guanine-rich oligonucleotide sequence (GROs) and monovalent cations in a growth buffer. AFM imaging of individual G-wire DNA strands requires many carefully monitored steps, including substrate preparation, G-wire concentration, adsorption onto substrate, rinsing, drying, appropriate selection/use of imaging probes, and dry atmosphere imaging conditions. Detailed step-wise instructions are provided. Key words: G-wire, Quadruplex DNA, Guanine-rich oligonucleotide, Self-assembly, Atomic force microscopy

1. Introduction G-DNA is a polymorphic family of four-stranded structures containing guanine tetrad motifs (1, 2) (see Fig. 1a). Guanine-rich oligonucleotides (GROs) that are self-complementary, as found in many telomeric (chromosome ends) repeat sequences (3, 4), form G-DNA in the presence of monovalent and/or divalent metal cations. The length and number of guanines and linker residues in GROs determine their diverse topologies. Quadruplex DNA has been constructed from mono, double, and quadruple strands of GROs, and are looped or linear, parallel or antiparallel (4), with a minimum of two base-stacked G-quartets (Fig. 1b). Naturally occurring hairpin structures, comprised of guanine quartets, are thought to play a role in telomerase activity that is essential for DNA replication (5). GROs can self-assemble in the

Giampaolo Zuccheri and Bruno Samorì (eds.), DNA Nanotechnology: Methods and Protocols, Methods in Molecular Biology, vol. 749, DOI 10.1007/978-1-61779-142-0_8, © Springer Science+Business Media, LLC 2011

105

106

Vesenka

Fig. 1. Quadruplex DNA is composed of G-quartets (a). The Tet1.5 monomer can form a dimer pair with a “closed,” “looped,” or “staggered” conformation as shown in (b). In either of the closed or looped conformations, no more growth of the G-wires can occur. In the staggered conformation, another dimer can attach to the G-wire ladder creating a succession of “sticky ends,” enabling multimers to assemble. The process is driven thermodynamically (14 ). Monomeric cation species, such as potassium or sodium, are known to stabilize the G-wires down the base-stacked core of the structure as seen in panel (b) (3, 4 ).

presence of monovalent cations to micrometer length linear chains, hence the term “G-wires.” The images shown in this work are from Tetrahymena thermophila with the oligonucleotide sequence of G4T2G4 (Tet1.5). Individual strands are easily imaged by atomic force microscopy (AFM) on the surface of mica because they are exceptionally stable to electrostatic collapse, unlike ­double-stranded DNA (see Fig.  2) (6). Long chain quadruplex DNA is of interest to nano- and biotechnological fields as ­templates for molecular wires (7) and as therapeutics (8). The procedure involved in successful growth of G-wire DNA involves starting with highly purified oligonucleotide sequences.

Preparation and Atomic Force Microscopy of Quadruplex DNA

b

K+ Mg2 G +

-

N

Nc -

-

G

M+ G G

107

-

-

Fig. 2. Electrostatic pinning (6 ) suggests that the greater internal attractive forces of G-wire DNA, comprised of guanine-quartet building blocks, four in a row, enable it to retain its solution-state and crystal-state structure when exposed to tether cations, here shown as magnesium. However, the stronger tethering force exerted on the unsupported phosphate backbone of duplex DNA, shown here as N–Nc (complementary) base pair, electrostatically pins the duplex DNA flat to the mica substrate.

These can either be purified from natural sources (9) or be ­synthetically constructed (10). The next important consideration is to include the appropriate monovalent cations in the buffer at sufficient oligonucleotide concentrations so that long G-wires are formed. Lastly, successful AFM imaging of G-wire DNA involves dilution (for individual strand observation), rapid adhesion onto smooth substrates, and thorough rinsing with deionized water to remove undesirable salt artifacts from the imaging process.

2. Materials With few exceptions (3), GROs self-assemble into G-wires in the presence of potassium or sodium. Quadruplex DNA formed in the presence of sodium tends to be interplanar or planar with the G-quartets, whereas those formed in the presence of potassium are almost exclusively interplanar (11, 12). Thus growth buffers must contain one of these two ions. Temperature seems to play little role in the growth or stability of many G-wires (13). The self-assembly process is concentration driven (14). Successfully grown G-wires require a growth concentration in the neighborhood of 100  mM, but must be diluted for AFM imaging of individual G-wires.

108

Vesenka

2.1. GRO Self-assembly

1. Growth buffer: 5 mM NaCl (or KCl), 10 mM MgCl2, 10 mM Tris–HCl, pH 7.5, and 1 mM spermidine (see Note 1). Though optimized for the Tet1.5 sequence, this growth buffer appears to work for many other naturally occurring GROs (J. Vesenka, unpublished data). 2. Obtain Tet1.5 from natural sources (9) or it can be synthetically made. For Tet1.5 (G4T2G4) sequence, 1OD = 1A260 of lyophilized G4T2G4 = 9.99  nmole. Dissolver this amount in 100 mL of growth buffer to provide a starting concentration of 100 mM, the minimum recommended for successful G-wire self-assembly (see Note 2).

2.2. Imaging Buffers

The presence of divalent cations, such as magnesium, improves the adsorption of duplex (15) and quadruplex (6) DNA onto the substrate of choice, mica. One consequence of the use of divalent cations to tether DNA onto mica is that it leads to collapse of duplex DNA, whereas the inherent stability of the quadruplex DNA helps to maintain its structure (Fig. 2). The growth buffer described previously will work satisfactorily as an imaging buffer. Salt artifacts can be further reduced in AFM images by substituting acetate (Ac) for chloride ions because of acetate’s greater volatility [e.g., 5 mM NaAc (or KAc), 10 mM MgAc2, and 10 mM Tris–Ac, pH 7.5].

3. Methods 3.1. GRO Self-assembly

No special temperature other than being in a liquid state is required for self-assembly of G-wires. After 24 h of incubation in the buffer described in Subheading  2.1, three samples were deposited and imaged on the surface of mica (procedure described shortly). At 100 mM concentration, the entire surface is coated with strands of DNA (Fig. 3a). At 10 mM concentration, the “DNA network” is

Fig. 3. Fresh, lyophilized Tet1.5 GRO incubated for 24 h in growth buffer. (a) 100 mM of incubated GRO adsorbed on mica for 10 min, rinsed, dried, and imaged. (b) 10 mM of incubated GRO adsorbed on mica for 10 min, rinsed, dried, and imaged. (c) 1 mM of incubated GRO adsorbed on mica for 10 min, rinsed, dried, and imaged. Note the transition from packed strands to a G-wire network to individual wires. Average length of individual wires in the image was 70 ± 30 nm and their average height was (equivalent diameter) 2.0 ± 0.1 nm.

Preparation and Atomic Force Microscopy of Quadruplex DNA

109

commonly observed (Fig. 3b). At 1 mM concentration, G-wires are observed with lengths in the 70 ± 30-nm range and 2 ± 0.1 nm in height (equivalent to diameter – Fig. 3c). If the concentration is decreased by another order of magnitude, typically no linear structures can be found, presumably because the G-wires have disassembled into constituent GROs. Low concentrations (1 mM) and longer adsorption times (at least 1  h) onto mica commonly lead to “auto-orientation” of quadruplex DNA with the hexagonal surface of mica (Fig.  4a) (16). Low concentrations and extremely long adsorption times (weeks) will lead to “rafting” of the G-wires (Fig. 4b, c). When growing G-wires at elevated temperature, e.g., 37°C, evaporation of buffer can present a problem. Evaporation can be remedied by injecting a layer of fresh mineral oil that will float over the

Fig. 4. A notable feature of the shorter segments of quadruplex DNA is their ability to “auto-orient” on the surface of mica (17 ). (a) After 1 h of incubation, the auto-orientation is easily observed. (b, c) Longer (weeks) incubation times lead to the self-assembly of G-wire “rafts,” still with noticeable auto-orientation. The rinse and blast drying do not appear to affect the orientation, presumably because of the negligible impact of surface tension on such small molecules.

110

Vesenka

GRO–buffer ­mixture (see Note 3). Long-term storage at elevated temperatures is not advisable as the samples appear to discolor, possibly due to impurities from the mineral oil interacting with the GROs–buffer. However, long-term storage at room temperature, 4°C, or freezing appears to have no impact on G-wire integrity. 3.2. AFM Sample Preparation and Imaging

1. Attach a mica disk (high-quality, 10 mm diameter disks were used in this work) to a ferrous disk (steel “puck”) using adhesive tabs (17). The metal disk can be secured to a magnet for the purposes of cleaving mica, rinsing, and drying. 2. Freshly cleave the mica substrate using transparent tape by firmly pressing the tape onto mica and pulling the tape off as if you were opening a hard cover book. Peeling mica off by rolling the tape will damage the mica. 3. Examine the fresh mica surface for mirror smoothness. This will aid in uniform spreading of the G-wire sample. Mica that appears “scratched” should be cleaved again or discarded. 4. Place 20  mL of sample onto the mica and check for uniform spreading of the solution over the disk. Uniform spreading is an indication of a clean substrate and will provide good G-wire adhesion. A sample that does not spread will result in poor AFM images. Repeat from step 2 if the sample does not spread. 5. Incubate on the mica for the desired length of time (from seconds to weeks). Do not let the sample dry, as dried buffer salts ruin AFM imaging (see Note 4). 6. Rinse with 1 ml of deionized water. This is most easily accomplished by direct deposition of the water onto the mica/puck attached to a secure magnet. Draining the sample is not necessary. 7. Immediately after rinsing, “blast dry” the sample with dry nitrogen at a gauge pressure of about 20  psi = 140  kPa. A nozzle connected to flexible Tygon™ tubing attached to a tank of dry nitrogen with an easy open valve works well. A 1  mL pipette tip with its end snipped off makes a good nozzle (see Note 5). 8. The sample should then be imaged immediately or stored in a dry environment until ready for imaging. Relative humidity above 25% will lead to migration of residual salts on the hygroscopic mica surface (18) and will ruin imaging. 9. To optimize dynamic atomic force microscopy resolution of the samples prepared above, four elements should be considered: A vibrationally, electronically, and thermally stable microscope; low humidity; sharp AFM probes; and slow scans with small RMS amplitudes. Recent advances in improving the stability of scanning probe microscopes have allowed for

Preparation and Atomic Force Microscopy of Quadruplex DNA

111

real-time corrections at the atomic scale (19). Until these processes are commercialized, high-resolution imaging requires active vibration isolation (20) and several hours of exercising the piezoelectric scanner. Active scanning at the desired scan range to work out piezoelectric hysteresis is microscope dependent. Some AFM systems allow this to be operator controlled (e.g., NDT), and other systems require “false engagement” (e.g., Digital Instruments/Veeco) to avoid tip contact (see Note 6). 10. Humidity and thermal equilibrium can be achieved by installing the sample and tip in the microscope surrounded by a plastic cover. Run a low-pressure stream (see Note 7) of dry helium through a hose into the cover and allow the microscope to equilibrate for several hours while electronic stabilization is being performed. The dry helium has an added effect of slightly improving the quality value of the resonance peak. 11. Sharp hydrophobic AFM probes are now available from numerous manufacturers. These include materials such as carbon nanotubes (21) and diamond-like carbon whiskers (22). Hydrophobicity of the probe reduces the likelihood of contamination from the G-wires and extends probe life. Follow the recommended manufacturer’s instruction on the use of these tips to optimize their imaging performance (23). In brief, the equipment should be well stabilized and the RMS oscillation of the cantilever should be around 1 nm to reduce ruinous impulse between the AFM tip and sample. Scan sizes should be small (about 250 nm or less) and scan speeds at this size slow (0.1 Hz). The scan speed can be proportionally increased with decrease in scan size. The importance of system stability becomes obvious as scanner drift can obscure the image. 12. After stabilization, adjust the scan size to “0” and adjust the set point while still scanning to the desired RMS value, and re-engage the microscope. When the microscope has automatically contacted the surface, the set point can be increased to back off the tip from the surface and the scan size restored to the desired value. When ready to image, decrease the set point manually to a value just below the automatic engagement value and capture the image.

4. Notes 1. The monovalent cation stabilizes the G-quartet. The magnesium ions and spermidine stabilize the G-wire phosphate backbone. Prepare buffer from fresh deionized water at 10× the

112

Vesenka

concentration described above and aliquot into sterile 1  ml Eppendorf tubes and freeze at −20°C for later use. 2. To estimate the quantity of single-stranded GROs needed in micrograms for different sequences, use the following formulas (24):

1 A 260 unit (= 1OD) of single-stranded DNA » 33 mg/mL



#(mg) ´ 3.0 (nmole/(mg)/N = #(nmole), where N is the number of bases in the oligonucleotide sequence. 3. When taking an aliquot from a concentrated GRO solution above room temperature with mineral oil on the top, the sample should first be allowed to cool to room temperature to avoid gas expansion (and incorrect volume measurements) in the pipette tip. The pipette tip can then be immersed through the oil layer with the plunger pre-depressed to a selected volume slightly greater than desired. After collecting the sample, wipe the outer edges of the pipette tip on wax film to remove excess oil. Deposit the aliquot on a fresh piece of wax film and use a fresh pipette tip to siphon off the desired volume of the aqueous droplet that remains, leaving behind any remaining mineral oil residue. This process can be used down to 1 mL effectively. 4. If the ambient humidity is low, evaporation will be easily noticed. At 10% relative humidity, a 20 mL sample will evaporate in less than 30  min. Long incubation times necessitate keeping the sample moist. This is most conveniently achieved by placing the freshly made sample on an elevated platform inside a Petri dish (e.g., 3 cm diameter) with 1 mL of deionized water on the bottom of the dish. Cover the dish and seal it with wax tape (e.g., Parafilm™) and secure in a safe place until you are ready to proceed to imaging. 5. A fast stream of nitrogen decreases the size of salt crystals, but can literally blow away a weakly anchored layer of mica. A clean white Styrofoam shipping box can help to monitor this procedure: examine the box after drying the sample to see if mica fragments are visible. If no DNA is found upon AFM imaging, the culprit may very well be that the sample layer was accidentally removed. In either case, the sample preparation must be repeated. 6. “False engagement” can be achieved on a Digital Instruments/ Veeco Multimode AFM by reducing the amplitude set point in Tappingmode™ to “0”. The software will cause the scanner to engage falsely and scan at the speed and size determined by the user. 7. A low stream of helium can be established by the “lip test.” Adjust the stream so that it just barely registers on whetted lips.

Preparation and Atomic Force Microscopy of Quadruplex DNA

113

References 1. Williamson, J.R., Raghuraman, M.K., and Cech, T.R. (1989) Monovalent cationinduced structure of telomeric DNA: the G-quartet model. Cell. 59, 871–880. 2. Williamson, J.R. (1993) G-Quartets in Biology: Reprise. Proc. Natl. Acad. Sci. USA. 90, 3124–3124. 3. Burge, S., Parkinson, G. N., Hazel, P., Todd, A. K., and Neidle, S. (2006) Quadruplex DNA: sequence, topology and structure Nucleic Acids Research, 34:19, 5402–5415. 4. Williamson, JR. (1994) G-quartet structures in telomeric DNA Annu Rev Biophys Biomol Struct. 23, 703–730. 5. Sen, D. and Gilbert W. (1992) Novel DNA superstructures formed by telomere-like oligomers. Biochem. 31, 65. 6. Muir, T., Morales, E., Root, J., Kumar, I., Garcia, B., Vellandi, C., Marsh, T., Henderson, E., and Vesenka, J. (1998) The morphology of duplex and quadruplex DNA on mica. J. Vac. Sci. Technol. A. 16, 1172–1177. 7. Weglarz, M., Fritzsche, W., Yerkes, S., Kmiec, E., and Vesenka, J. (2008) “Analysis of G5x Quadruplex DNA”, DNA-Based Nano-Scale Integration, AIP Conference Proceedings 1062, 123–128. 8. Borman, S., (2007) Ascent Of Quadruplexes: Nucleic acid structures become promising drug targets Chem. & Eng. News 85, 22, 12–17. 9. Marsh, T.C., Vesenka, J., and Henderson, E., (1995) A new DNA nanostructure, the G-wire, imaged by scanning probe microscopy, Nucleic Acids Res. 23:696–700. 10. Vesenka, J., Vellandi, C., Kumar, I., Marsh, T., and Henderson, E., (1998) The diameter of duplex and quadruplex DNA measured by Scanning Probe Microscopy. Scanning Microscopy 12:2, 329–342. 11. Howard, F. B., Frazier, J. and Miles, H. T. (1977) Stable and metastable forms of poly(G). Biopolymers, 16, 791–809. 12. Dingley, A. J., Peterson, R. D., Grzesiek, S. and Feigon, J. (2005) Characterization of the cation and temperature dependence of DNA quadruplex hydrogen bond properties using high-resolution NMR. J. Amer. Chem. Soc., 127, 14466–14472. Sci. USA, 102, 634–639.

13. Schwartz, T.R., Vasta, C.A. Bauer, T.L. Parekh-Olmedo, H., and Kmiec, E., (2008) G-Rich Oligonucleotides Alter Cell Cycle Progression and Induce Apoptosis Specifically in OE19 Esophageal Tumor Cells, Oligonucleotides 18, 51–63. 14. Marsh, T., and Vesenka, J. (2007) Properties of G-Wire DNA. Nano and Molecular Electronics Handbook, Sergy Lyshevski ed., CRC Press, New York, 13, 1–15. 15. Vesenka, J., Tang, C. L., Guthold, M., Keller, D., Delaine, E., and Bustamante, C. (1992) A substrate preparation for imaging biomolecules with the scanning force microscope. Ultramicroscopy, 42–44, 1243–1249. 16. Vesenka, J., Bagg, D., Wolff, A., Reichert, A., and Fritzsche, W., (2007) Auto-Orientation of G-wire DNA on Mica. Colloids and Surfaces B: Biointerfaces, 58, pp. 256–263. 17. E.g. (this is NOT a product endorsement) all supplies from http://www.tedpella.com/. 18. Vesenka, J., Manne, S., Yang, G., Bustamante, C., and Henderson, E., (1993) Humidity effects on atomic force microscopy of goldlabeled DNA on mica. Scanning Microscopy, 7, 781–788. 19. Perkins, T., King, G., Carter, A. and Churnside, A., (2008) Ultrastable atomic force microscopy: atomic-scale stability and registration in ambient conditions. AFMBiomed Conference, Monterey CA, October 15–18. 20. An animated summary can be found at http:// physics-animations.com/Physics/English/ spri_txt.htm. 21. Hall, A., Matthews, W. G., Superfine, R., Falvo, M. R., and Washburn, S., (2003) Simple and efficient method for carbon nanotube attachment to scanning probes and other substrates. Appl. Phys. Lett. 82, 2506. 22. Klinov, D., and Magonov, S., (2004) True molecular resolution in tapping-mode atomic force microscopy with high-resolution probes. Appl. Phys. Lett. 84, 2697. 23. E.g. (this is NOT a product endorsement) http://www.spmtips.com/howto/res/hr 24. E.g. (this is NOT a product endorsement) http://www.biosyn.com.

Chapter 9 Synthesis of Long DNA-Based Nanowires Alexander Kotlyar Abstract Here we describe novel procedures for production of DNA-based nanowires. This include synthesis and characterization of the one-to-one double-helical complex of poly(dG)–poly(dC), triple-helical poly(dG)– poly(dG)–poly(dC) and G4-DNA, which is a quadruple-helical form of DNA. All these types of DNAbased molecules were synthesized enzymatically using Klenow exo− fragment of DNA Polymerase I. All the above types of nanowires are characterized by a narrow-size distribution of molecules. The contour length of the molecules can be varied from tens to hundreds of nanometers. These structures possess improved conductive and mechanical properties with respect to a canonical random-sequenced DNA and can possibly be used as wire-like conducting or semiconducting nanostructures in the field of nanoelectronics. Key words: DNA nanowires, Enzymatic synthesis, Klenow exo−, Poly(dG)–poly(dC), G4-DNA, Triplex DNA

1. Introduction The DNA molecule is an attractive candidate to wire electrons over long molecular distances. Charge migration along DNA molecules has been the subject of scientific interest for many years. It is currently accepted by the scientific community that a native, random sequence DNA is not a good electrical molecular wire, due to its apparent poor intrinsic conductivity. It has been demonstrated that uniform DNA comprising repeating sequences improves conduction properties. Recent experimental demonstration of the conducting behavior in short poly(dG)–poly(dC) DNA oligomers (1, 2) and the results of theoretical calculations show that poly(dG)–poly(dC), a homopolymer consisting of a pair of poly(dC) and poly(dG) chains, exhibits better conductance than poly(dA)–poly(dT) homopolymer (3). This is mainly due to the fact that poly(dG)–poly(dC) provides better conditions

Giampaolo Zuccheri and Bruno Samorì (eds.), DNA Nanotechnology: Methods and Protocols, Methods in Molecular Biology, vol. 749, DOI 10.1007/978-1-61779-142-0_9, © Springer Science+Business Media, LLC 2011

115

116

Kotlyar

for p overlap compared to poly(dA)–poly(dT). In addition, ­guanine, which present in a high quantity in the former DNA is ­characterized by lowest ionization potential among DNA bases, thus promoting charge migration through the DNA (4). These facts emphasize the importance of guanine-rich DNA-based ­molecules as possible candidates for molecular electronics applications. The following guanine-rich DNA-based structures, whose syntheses is described here, may offer the desired electrical conduction properties (1) double-stranded poly(dG)–poly(dC), (2) triple-stranded poly(dG-dG)–poly(dC), and (3) four-stranded G4-DNA. A poly(dG)–poly(dC) is a double-stranded deoxyribopolynucleotide polymer, which consists of a pair of antiparallel poly(dC) and poly(dG) homopolymers. Commercial preparations of the DNA are available and were used by researchers in electrical conductivity studies (5, 6). We have demonstrated (7), however, that commercial preparations of poly(dG)–poly(dC) consists of long continuous poly(dC)-strand and relatively short poly(G) fragments, 500–1,500 bases long, associated with the C-strand, but not covalently connected to each other. The presence of the G-strand breaks along poly(dG)–poly(dC) must strongly reduce the ability of the polymer to conduct current and strongly limits the use of the ­molecules in nanoelectronics. The molecules prepared by our technique (7) lack the above disadvantages. The enzymatic synthesis, conducted as described below, yielded doublestranded poly(dG)–poly(dC) characterized by a well-defined length (up to 10 kb) and narrow-size distribution of molecules. The synthesized molecules composed of continuous dG- and dChomopolymers of equal length lacking strand nicks. In addition, the poly(dG)–poly(dC) may comprise a functional group attached to 5¢ ends of either one or both strands composing the DNA (7). The functional group may be a particular sequence of singlestranded DNA, fluorescent labels, thiol-groups, biotin moieties, and other groups. Thiols are known to interact specifically with gold (8, 9). Introduction of SH-groups at the ends of DNA was used to anchor the DNA fragments to flat gold surfaces (10). The ability to attach SH-groups to the 5¢ ends of the strands thus provides a tool for the selective binding of long poly(dG)–poly(dC) polymers to gold surfaces and gold nanoelectrodes. This property is especially useful for application of the polymer in nanoelectronics. Triple-stranded DNA structures have been a subject of research in the past 50 years ( for review, see Refs. 11–13). Most of these structures are composed of tens of triads. Long triplestranded DNA, poly(dG-dG)–poly(dC) have been reported by us only recently (14). A poly(dG-dG)–poly(dC) is an intramolecular triplex, composed of continuous dG- and dC-homopolymers. The length of the dG-homopolymer is twice the length of the

Synthesis of Long DNA-Based Nanowires

117

poly(dC), which enables the former strand to fold back to pair with the adjacent of poly(dG)–poly(dC) duplex. The length of the latter triplex structures can be varied from ten to hundreds of nanometers. We were able to obtain the triplex nanostructures characterized by very narrow-size distribution by applying the enzymatic method described below. As in the case of poly(dG)– poly(dC) molecules, various functional group including fluorescent labels, thiol-groups can be attached to 5¢ ends of either one or both strands composing the triplex (14). We have demonstrated that the triplex molecules are stiffer and more resistant to mechanical deformation compared to random sequence DNA and poly(dG)–poly(dC) (14). This property, together with the ability to functionalize the synthesized triplex molecules is essential for their application as elements in nanodevices. G4-DNA wire is a very stable molecule made of consecutive stacked arrangement of 4 guanine (G) bases (tetrads – a tetrad consists of 4 G bases). It is known for decades that G-rich DNA sequences containing runs of guanines (dG) can form G-quadruplex structures ( for review see Refs. 15–17). These structures, commonly named G4-DNA, are comprised of stacked tetrads; each of the tetrads arises from the planar association of four guanines by Hoogsteen hydrogen bonding. Most of the studies have been performed using short (16–32 bases) G-rich telomeric oligonucleotides (18, 19). Short G-rich oligonucleotides were shown to assemble spontaneously into long molecular wires in the presence of proper monovalent cations (20, 21). These wires are very polymorphic, and constructed of short oligomers, resulting in nonuniform polymers with gaps (noncovalently bonded backbone) between G-rich oligonucleotide fragments along the formed wires (20, 21). Guanine tetrads were proposed as building blocks of molecular nanodevices (16, 17, 22). However, the above wires, formed by many short DNA segments, and containing many nicks are probably not good candidates for application as molecular nanowires. For the utilization of G4-DNA in nanoelectronics, long, persistent, homogenous populations of molecules are required. Only recently, we have reported a method (described below) for synthesis of novel long (hundreds of ­nanometers) continuous G-based nanostructures, composed of ­hundreds of stacked tetrads (23). These nanostructures are characterized by a narrow length distribution and contain no gaps in their backbone. We have also demonstrated that these wires are characterized by higher stability, resistance to heat treatment and higher charge polarizability, as compared to double-stranded DNA (24). These properties make these structures very promising for nanoelectronic applications. Enzymatic synthesis of the long, di, tri, and tetra-stranded G-rich DNA-based nanostructures is described in detail below.

 

118

Kotlyar

2. Materials 2.1. Preparation of (dG)12–(dC)12 Template–Primer

1. 0.1 M NaOH solution: 4 g of NaOH. Add 1 L of fresh water purified by a Seralpur Pro 90 CN system (Merck Belgolabo, Overijse, Belgium), deionized water and filtered through 0.22  mm Millipore Express PLUS membrane filter, deionized/filtered water. 2. 0.1  M NaOH containing 10% acetonitrile: 4  g of NaOH. Add 0.9 L of deionized/filtered water. Add 100 mL of acetonitrile (Bio Lab, HPLC-S Gradient grade). 3. 1  M phosphate buffer (pH 7.5): 136  g of KH2PO4. Add 700 mL of deionized water; adjust the pH to 7.5 with 1 M KOH and add deionized water to 1  L. Filter the solution through 0.22 mm Millipore Express PLUS membrane filter. 4. 20  mM Phosphate buffer (pH 7.5) containing 10% acetonitrile: to 880 mL of deionized water add 20 mL of 1 M Phosphate buffer (pH 7.5) and filter the solution through 0.22  mm Millipore Express PLUS membrane filter. Add 100 mL of acetonitrile (Bio Lab, HPLC-S Gradient grade). 5. 250  mM Phosphate buffer (pH 7.5) containing 10% acetonitrile: to 650 mL of deionized water add 250 mL of 1 M Phosphate buffer (pH 7.5) and filter the solution through 0.22  mm Millipore Express PLUS membrane filter. Add 100 mL of acetonitrile (Bio Lab, HPLC-S Gradient grade). 6. Alkaline 1 M NaCl solution containing acetonitrile: 58 g of NaCl (Merck). Add 0.9 L of 0.1 M NaOH and filter the solution through 0.22  mm Millipore Express PLUS membrane filter. Add 100 mL of acetonitrile (Bio Lab, HPLC-S Gradient grade). 7. Glacial acetic acid 100%. 8. 1 M Tris–acetate (pH 7.5): 121 g of Tris-base. Add 800 mL of deionized water; adjust the pH to 7.5 with glacial acetic acid and add deionized water to 1  L. Filter the solution through 0.22 mm Millipore Express PLUS membrane filter. 9. 2 mM Tris–acetate (pH 7.5): to 998 mL of deionized/­filtered water add 2 mL of 1 M Tris–acetate (pH 7.5). 10. 20  mM Tris–acetate (pH 7.5): to 980  mL of deionized/­ filtered water add 20 mL of 1 M Tris–acetate (pH 7.5). 11. (dG)12 oligonucleotide from Alpha DNA (Montreal, Canada). 12. (dC)12 oligonucleotide from Alpha DNA (Montreal, Canada). 13. Dialysis tubing 10 mm (Sigma). Treat the tubing as follows: wash with running tap water for 3 h; treat with 0.3% (w/v) solution of sodium sulfide at 80°C for 1 min; wash with tap

Synthesis of Long DNA-Based Nanowires

119

water for 2–5 min at 60°C; treat with 0.2% (v/v) solution of sulfuric acid for 10 min at room temperature; wash with tap water for 10–15 min. Store the tubing in 25% ethanol at 4°C. Rinse the tubing with running deionized/filtered water before use. 14. Ion-exchange PolyWax LP column (4.6 × 200  mm, 5  mm, 1,000 Å) (Western Analytical Products). 15. Sephadex NAP-25 DNA-Grade column (15 × 50  mm) (GE Healthcare). 16. Ion-exchange HiTrap Q HP column (1 mL) (GE Healthcare). 17. Agilent 1100 HPLC system with a photodiode array detector. 18. Eppendorf table centrifuge (model 5424). 19. Laboratory Freeze Dryer Christ Alpha 1–4 (Osterode am Harz, Germany). 2.2. Preparation of Thiol-End-Labeled (dG)12–(dC)12 Template–Primer [SH-(dG)12–(dC)12-SH]

1. 0.1  M NaOH containing 10% acetonitrile: 4  g of NaOH. Add 0.9 L of deionized/filtered water. Add 100 mL of acetonitrile (Bio Lab, HPLC-S Gradient grade). 2. 5  M HCl solution: to 215  mL of deionized/filtered water add 285 mL of 32% HCL (Merck). 3. Alkaline 1 M NaCl solution containing acetonitrile: 58 g of NaCl. Add 0.9  L of 0.1  M NaOH and filter the solution through 0.22 mm Millipore Express PLUS membrane filter. Add 100  mL of acetonitrile (Bio Lab, HPLC-S Gradient grade). 4. 1 M Tris–acetate (pH 7.5): 121 g of Tris-base. Add 800 mL of deionized water; adjust the pH to 7.5 with glacial acetic acid and add deionized water to 1  L. Filter the solution through 0.22 mm Millipore Express PLUS membrane filter. 5. 2 mM Tris–acetate (pH 7.5): to 998 mL of deionized/­filtered water add 2 mL of 1 M Tris–acetate (pH 7.5). 6. 20  mM Tris–acetate (pH 7.5): to 980  mL of deionized/­ filtered water add 20 mL of 1 M Tris–acetate (pH 7.5). 7. 0.4  M dl-dithiothreitol (DTT). 15.4  mg of DTT. Add 0.25 mL deionized/filtered water. Store at −18°C. 8. 5¢-(6-Mercapto-1-hexyl-phosphoric acid ester) of (dG)12 oligonucleotide, SH-(dG)12, from Alpha DNA (Montreal, Canada). 9. 5¢-(6-Mercapto-1-hexyl-phosphoric acid ester) of (dC)12 oligonucleotide, SH-(dC)12, from Alpha DNA (Montreal, Canada). 10. Dialysis tubing 10 mm (Sigma). Treat the tubing as follows: wash with running tap water for 3 h; treat with 0.3% (w/v)

120

Kotlyar

solution of sodium sulfide at 80°C for 1 min; wash with tap water for 2–5 min at 60°C; treat with 0.2% (v/v) solution of sulfuric acid for 10 min at room temperature; wash with tap water for 10–15 min. Store the tubing in 25% ethanol at 4°C. Rinse the tubing with running water and then with deionized/filtered water before use. 11. Ion-exchange PolyWax LP column (4.6 × 200  mm, 5  mm, 1,000 Å), Western Analytical Products. 12. Sephadex NAP-25 DNA-Grade (15 × 50 mm) (GE Healthcare).

prepacked

column

13. Ion-exchange HiTrap Q HP column (1 mL) (GE Healthcare). 14. Agilent 1100 HPLC system with a photodiode array detector. 15. Eppendorf table centrifuge (model 5424). 16. Laboratory Freeze Dryer Christ Alpha 1–4 (Osterode am Harz, Germany). 2.3. Synthesis of Poly(dG)–Poly(dC) 2.3.1. Enzymatic Synthesis of Poly(dG)–Poly(dC)

1. 5 M KOH solution: 140 g of KOH. Add 0.5 L of deionized water. 2. 1  M Phosphate buffer (pH 7.5): 136  g of KH2PO4. Add 700 mL of deionized water; adjust the pH to 7.5 with 5 M KOH and add deionized water to 1  L. Filter the solution through 0.22 mm Millipore Express PLUS membrane filter. 3. 1 M MgCl2: 20.3 g of MgCl2·6H2O. Add 100 mL of deionized water. Filter the solution through 0.22  mm Millipore Express PLUS membrane filter. 4. 1 M EDTA: 29.2 g of Titriplex II (Merck). Add 100 mL of deionized water. Filter the solution through 0.22  mm Millipore Express PLUS membrane filter. 5. 0.4  M dl-dithiothreitol (DTT). 15.4  mg of DTT. Add 0.25 mL of deionized/filtered water. Store at −18°C. 6. 100 mM dCTP. Dissolve 23.2 mg of dCTP (Sigma) in 0.5 mL of deionized/filtered H2O. Store at −18°C. 7. 100 mM dGTP. Dissolve 25 mg of dGTP (Sigma) in 0.5 mL of deionized/filtered water. Store at −18°C. 8. 10  mM (dG)12–(dC)12 prepared as described below (see Subheading 3.1). 9. Klenow exo− (Klenow fragment of Escherichia coli DNA polymerase I, lacking the 3¢ → 5¢ exonuclease activity), 5  U/mL enzyme solution in glycerol from Fermentas (Lithuania). Store the solution at −18°C. 10. Dry bath incubator (MRC, Israel).

Synthesis of Long DNA-Based Nanowires 2.3.2. HPLC Purification of Synthesized Poly(dG)–Poly(dC)

121

1. 1  M acetic acid solution: to 470  mL of deionized/filtered water add 30 mL of 100% acetic acid (Merck). 2. 20  mM Tris–acetate (pH 8.0): 2.42  g of Tris-base (Fluka). Add 800 mL of deionized water; adjust the pH to 8.0 with 1  M acetic acid and add deionized water to 1  L. Filter the solution through 0.22 mm Millipore Express PLUS membrane filter. 3. TSK-gel G-DNA-PW HPLC column (7.8 × 300  mm) (TosoHaas, Japan). 4. Agilent 1100 HPLC system with a photodiode array detector.

2.4. Synthesis of Thiol-End-Labeled Poly(dG)–Poly(dC), SH-Poly(dG)– Poly(dC)-SH 2.4.1. Enzymatic Synthesis of SH-Poly(dG)–Poly(dC)-SH

1. 5 M KOH solution: 140 g of KOH. Add 0.5 L of deionized water. 2. 1  M Phosphate buffer (pH 7.5): 136  g of KH2PO4. Add 700 mL of deionized water; adjust the pH to 7.5 with 5 M KOH and add deionized water to 1  L. Filter the solution through 0.22 mm Millipore Express PLUS membrane filter. 3. 1 M MgCl2: 20.3 g of MgCl2·6H2O. Add 100 mL of deionized water. Filter the solution through 0.22  mm Millipore Express PLUS membrane filter. 4. 1 M EDTA: 29.2 g of Titriplex II (Merck). Add 100 mL of deionized water. Filter the solution through 0.22  mm Millipore Express PLUS membrane filter. 5. 0.4  M dl-dithiothreitol (DTT). 15.4  mg of DTT (Sigma). Add 0.25 mL deionized/filtered water. Store at −18°C. 6. 100 mM dCTP. Dissolve 23.2 mg of dCTP (Sigma) in 0.5 mL of deionized/filtered H2O. Store at −18°C. 7. 100 mM dGTP. Dissolve 25 mg of dGTP (Sigma) in 0.5 mL of deionized/filtered water. Store at −18°C. 8. 10  mM SH-(dG)12–(dC)12-SH prepared as described below (see Subheading 3.2). 9. Klenow exo− (Klenow fragment of E. coli DNA polymerase I, lacking the 3¢ → 5¢ exonuclease activity), 5  U/mL enzyme solution in glycerol from Fermentas (Lithuania). Store the solution at −18°C. 10. Dry bath incubator (MRC, Israel).

2.4.2. HPLC Purification of Synthesized SH-Poly(dG)– Poly(dC)-SH

1. Glacial acetic acid 100%. 2. 1 M Tris–acetate (pH 7.5): 121 g of Tris-base (Fluka). Add 800 mL of deionized water; adjust the pH to 7.5 with glacial acetic acid and add deionized water to 1 L. Filter the solution through 0.22 mm Millipore Express PLUS membrane filter. 3. 2  mM Tris–acetate (pH 7.5): to 998  mL of deionized/ filtered water add 2 mL of 1 M Tris–acetate (pH 7.5).

122

Kotlyar

4. 20 mM Tris–acetate (pH 7.5): to 980 mL of deionized water add 20 mL of 1 M Tris–acetate (pH 7.5). 5. TSK-gel G-5000-PW HPLC column (7.8 × 300  mm), TosoHaas, Japan. 6. Agilent 1100 HPLC system with a photodiode array detector. 2.5. Synthesis of Poly(dG-dG)–Poly(dC) Triplex 2.5.1. Poly(dG-dG)– Poly(dC) Triplex Synthesis

1. 1  M KOH solution: 56  g of KOH. Add 1  L of deionized water. Filter the solution through 0.22 mm Millipore Express PLUS membrane filter. 2. 1 M Phosphate buffer (pH 7.5): 136 g of KH2PO4 (Merck). Add 700 mL of deionized water; adjust the pH to 7.5 with 1 M KOH and add deionized water to 1 L. Filter the solution through 0.22 mm Millipore Express PLUS membrane filter. 3. 1 M MgCl2: 20.3 g of MgCl2·6H2O. Add 100 mL of deionized water. Filter the solution through 0.22  mm Millipore Express PLUS membrane filter. 4. 1 M EDTA: 29.2 g of Titriplex II (Merck). Add 100 mL of deionized water. Filter the solution through 0.22  mm Millipore Express PLUS membrane filter. 5. 0.4  M dl-dithiothreitol (DTT). 15.4  mg of DTT. Add 0.25 mL deionized/filtered water. Store at −18°C. 6. 100 mM dGTP. Dissolve 25 mg of dGTP (Sigma) in 0.5 mL of deionized/filtered water. Store at −18°C. 7. 1 mM (in base pairs) 500–1,000 base pairs poly(dG)–poly(dC) prepared as described in Subheading 3.3. 8. Klenow exo− (Klenow fragment of E. coli DNA polymerase I, lacking the 3¢ → 5¢ exonuclease activity), 5  U/mL enzyme solution in glycerol from Fermentas (Lithuania). Store the solution at −18°C. 9. Dry bath incubator (MRC, Israel).

2.5.2. HPLC Purification of Synthesized Poly(dG-dG)–Poly(dC)

1. 1  M acetic acid solution: to 470  mL of deionized/filtered water add 30 mL of 100% glacial acetic acid (Merck). 2. 20  mM Tris–acetate (pH 7.5): 2.42  g of Tris-base. Add 800 mL of deionized water; adjust the pH to 7.5 with 1 M acetic acid and add deionized water to 1 L. Filter the solution through 0.22 mm Millipore Express PLUS membrane filter. 3. TSK-gel G-DNA-PW HPLC column (7.8 × 300  mm), TosoHaas, Japan. 4. Agilent 1100 HPLC system with a photodiode array detector.

123

Synthesis of Long DNA-Based Nanowires

2.6. Synthesis of G4 (Quadruple)-DNA 2.6.1. Purification of (dC)20

1. 0.1 M NaOH solution: 4 g of NaOH. Add 1 L of deionized water. Filter the solution through 0.22 mm Millipore Express PLUS membrane filter. 2. Glacial acetic acid 100% (Merck). 3. 1  M Phosphate buffer (pH 7.5): 136  g of KH2PO4. Add 700 mL of deionized water; adjust the pH to 7.5 with 1 M KOH and add deionized water to 1  L. Filter the solution through 0.22 mm Millipore Express PLUS membrane filter. 4. 20  mM Phosphate buffer (pH 7.5) containing 10% acetonitrile; to 880 mL of deionized water add 20 mL of 1 M Phosphate buffer (pH 7.5) and filter the solution through 0.22  mm Millipore Express PLUS membrane filter. Add 100 mL of acetonitrile (Bio Lab, HPLC-S Gradient grade). 5. 0.5 M Phosphate buffer (pH 7.5) containing 10% acetonitrile; to 0.4 L of deionized water add 0.5 L of 1 M Phosphate buffer (pH 7.5) and filter the solution through 0.22 mm Millipore Express PLUS membrane filter. Add 100 mL of acetonitrile (Bio Lab, HPLC-S Gradient grade). 6. 1 M Tris–acetate (pH 7.5): 121 g of Tris-base (Fluka). Add 800 mL of deionized water; adjust the pH to 7.5 with glacial acetic acid and add deionized water to 1 L. Filter the solution through 0.22 mm Millipore Express PLUS membrane filter. 7. Tris–acetate (pH 7.5): to 998 mL of deionized/filtered water add 2 mL of 1 M Tris–acetate (pH 7.5). 8. 10  mM (dG)12–(dC)12 Subheading 3.1).

prepared

as

described

(see

9. (dC)20 oligonucleotide from Alpha DNA (Montreal, Canada). 10. Dialysis tubing 10 mm (Sigma). Treat the tubing as follows: wash with running tap water for 3 h; treat with 0.3% (w/v) solution of sodium sulfide at 80°C for 1 min; wash with tap water for 2–5 min at 60°C; treat with 0.2% (v/v) solution of sulfuric acid for 10 min at room temperature; wash with tap water for 10–15 min. Store the tubing in 25% Ethanol at 4°C. Rinse the tubing with running deionized/filtered water before use. 11. Ion-exchange PolyWax LP column (4.6 × 200  mm, 5  mm, 1,000 Å), Western Analytical Products. 12. Sephadex NAP-25 DNA-Grade (15 × 50 mm), GE Healthcare.

prepacked

column

13. Ion-exchange HiTrap Q HP column (1 mL), GE Healthcare. 14. Agilent 1100 HPLC system with a photodiode array detector. 15. Eppendorf table centrifuge (model 5424). 16. Laboratory Freeze Dryer Christ Alpha 1–4 (Osterode am Harz, Germany).

124

Kotlyar

2.6.2. Poly(dG)–n(dC)20 Synthesis

1. 1  M KOH solution: 56  g of KOH. Add 1  L of deionized water. Filter the solution through 0.22 mm Millipore Express PLUS membrane filter. 2. 1  M Phosphate buffer (pH 7.5): 136  g of KH2PO4. Add 700 mL of deionized water; adjust the pH to 7.5 with 1 M KOH and add deionized water to 1  L. Filter the solution through 0.22 mm Millipore Express PLUS membrane filter. 3. 20 mM Phosphate buffer (pH 7.5): to 98 mL of deionized water add 2 mL of 1 M Phosphate buffer (pH 7.5). Filter the solution through 0.22  mm Millipore Express PLUS membrane filter. 4. 1 M MgCl2: 20.3 g of MgCl2·6H2O. Add 100 mL of deionized water. Filter the solution through 0.22  mm Millipore Express PLUS membrane filter. 5. 1 M EDTA: 29.2 g of Titriplex II (Merck). Add 100 mL of deionized water. Filter the solution through 0.22  mm Millipore Express PLUS membrane filter. 6. 0.4  M dl-dithiothreitol (DTT). 15.4  mg of DTT. Add 0.25 mL deionized/filtered water. Store at −18°C. 7. 100 mM dGTP. Dissolve 25 mg of dGTP (Sigma) in 0.5 mL of deionized/filtered H2O. Store at −18°C. 8. 10 mM (dG)12–(dC)12 template/primer prepared as described below (see Subheading 3.1). 9. HPLC-purified (dC)20 prepared as described below (see Subheading 3.6.1). 10. Klenow exo− (Klenow fragment of E. coli DNA polymerase I, lacking the 3¢ → 5¢ exonuclease activity), 5  U/mL enzyme solution in glycerol from Fermentas (Lithuania). Store the solution at −18°C. 11. Dry bath incubator (MRC, Israel).

2.6.3. HPLC Purification of Synthesized Poly(dG)–n(dC)20

1. 1  M acetic acid solution: to 470  mL of deionized/filtered water add 30 mL of 100% glacial acetic acid. 2. 20  mM Tris–acetate (pH 7.5): 2.42  g of Tris-base. Add 800 mL of deionized water; adjust the pH to 7.5 with 1 M acetic acid and add deionized water to 1 L. Filter the solution through 0.22 mm Millipore Express PLUS membrane filter. 3. TSK-gel DNA-G-DNA PW HPLC column (7.8 × 300 mm) from TosoHaas, Japan. 4. Agilent 1100 HPLC system with a photodiode array detector.

Synthesis of Long DNA-Based Nanowires 2.6.4. Preparation of Poly(dG) Strands

125

1. 1 M NaOH solution: 40 g of NaOH. Add 1 L of deionized water. Filter the solution through 0.22 mm Millipore Express PLUS membrane filter. 2. 0.1  M NaOH: to 900  mL of deionized/filtered water add 100 mL of 1 M NaOH. 3. TSK-gel DNA-G-DNA PW HPLC column (7.8 × 300 mm) from TosoHaas, Japan. 4. Agilent 1100 HPLC system with a photodiode array detector.

2.6.5. Preparation of G4-DNA

1. 1  M acetic acid solution: to 470  mL of deionized/filtered water add 30 mL of 100% glacial acetic acid. 2. 2  mM Tris–acetate (pH 8.0): 242  mg of Tris-base. Add 900 mL of deionized water; adjust the pH to 8.0 with 1 M acetic acid and add deionized water to 1 L. Filter the solution through 0.22 mm Millipore Express PLUS membrane filter. 3. Sephadex G-25 (NAP-5) prepacked DNA-Grade column (10 × 30 mm), GE Healthcare.

3. Methods 3.1. Preparation of (dG)12–(dC)12 Template–Primer 3.1.1. HPLC Purification of (dC)12

Complete purification of (dG)12 and (dC)12 oligonucleotides comprising a (dG)12–(dC)12 template–primer from shorter or longer oligonucleotides is required (see Notes 1 and 2). 1. Transfer ~1 mg of a dry oligonucleotide powder to 1.5 mL plastic tube. 2. Add 1 mL of deionized/filtered water. 3. Shake the sample and vortex vigorously for 2 min; incubate at room temperature for 30 min and vortex again. 4. Centrifuge the sample for 2  min at 5,000 × g at room temperature in order to get rid of insoluble compounds that might be present in the oligonucleotide preparation. 5. Transfer the entire supernatant to a new 1.5  mL plastic tube. 6. Connect an ion-exchange PolyWax LP column to the HPLC system. 7. Equilibrate the column with 50  mL of 20  mM Phosphate buffer (pH 7.5) containing 10% acetonitrile at a flow rate of 0.8 mL/min at room temperature. 8. Load 150 mL of the oligonucleotide sample at a flow rate of 0.8 mL/min. Do not overload the column; the large sample volume can significantly reduce the separation efficiency.

126

Kotlyar

Fig. 1. Purification of (dC)12 on a Poly WAX 300 Å column.

9. Elute the oligonucleotide in 10% acetonitrile with a linear Phosphate buffer gradient from 0.02 to 0.25 M for 30 min at a flow rate of 0.8 mL/min at room temperature. Monitor the elution by measuring absorbance at 260 nm. The elution profile is shown in Fig. 1. 10. Collect the fraction containing (dC)12 eluted between 33.5 and 35 min (as indicated by the arrows in Fig. 1). Total volume of the fraction should be ~1.0 mL. 11. Equilibrate the Sephadex G-25 DNA-Grade column with 30 mL of 2 mM Tris–acetate (pH 7.5) at room temperature. 12. Load 1 mL of the oligonucleotide sample obtained from the ion-exchange column (see step 10). Allow the sample to enter the column completely. Add 2.0 mL of Tris–acetate (pH 7.5). Allow the buffer to enter the column. 13. Place 2.0 mL plastic tube under the column; add 1.5 mL of 2 mM Tris–acetate (pH 7.5) and collect the eluant. 14. Transfer the sample into two 1.5 mL plastic tubes (1.0 mL per tube). 15. Freeze the sample in a dry ice/ethanol bath and lyophilize it to dryness. It takes approximately 15  h to completely lyophilize the sample. 16. Store the dry sample at −18°C. 3.1.2. HPLC Purification of (dG)12

1. Transfer ~1  mg of a dry oligonucleotide powder to plastic 1.5 mL plastic tube. 2. Add 1 mL of 0.1 M NaOH. 3. Shake the sample and vortex vigorously for 2 min.

Synthesis of Long DNA-Based Nanowires

127

Fig. 2. Purification of (dG)12 on a HiTrap Q HP column.

4. Centrifuge the sample for 2  min at 5,000 × g at room temperature in order to get rid of insoluble compounds that might be present in the oligonucleotide preparation. 5. Transfer the entire supernatant to a new 1.5  mL plastic tube. 6. Connect a HiTrap Q HP column to the HPLC system. 7. Equilibrate the column with 0.1  M NaOH containing 10% acetonitrile at a flow rate of 0.7  mL/min at room temperature. 8. Load 150 mL of the oligonucleotide sample at a flow rate of 0.7 mL/min. 9. Elute the oligonucleotide in 0.1  M NaOH containing 10% acetonitrile with a linear NaCl gradient from 0.5 to 1 M for 60 min at a flow rate of 0.7 mL/min at room temperature. Monitor the elution by measuring absorbance at 260  nm. The elution profile is shown in Fig. 2. 10. Collect the fraction eluted between 35 and 37 min (as indicated by the arrows in Fig. 2). Total volume of the fraction should be ~1.5 mL. 11. Equilibrate Sephadex G-25 DNA-Grade column with 30 mL of 2 mM Tris–acetate (pH 7.5) at room temperature. 12. Load 1.5  mL of the oligonucleotide sample obtained from the ion-exchange column (see step 10). Allow the sample to enter the column completely. Add 1.5  mL of 2  mM Tris– acetate (pH 7.5) buffer. Allow the buffer to enter the column. 13. Place 2.0  mL plastic tube under the column; add 2  mL of 2 mM Tris–acetate (pH 7.5) buffer and collect the eluant.

128

Kotlyar

14. Transfer the solution into two 1.5 mL plastic tubes (1.0 mL per tube). 15. Freeze the sample in a dry ice/ethanol bath and lyophilize it to dryness. It takes approximately 15  h to completely lyophilize the sample. 16. Store the dry sample at −18°C. 3.1.3. Annealing of Purified (dC)12 and (dG)12

1. Dissolve HPLC-purified (dC)12 obtained as described in Subheading 3.1.1 in 200 mL of 0.1 M NaOH. 2. Withdraw 10 mL from the sample and add to a quartz cuvette filled with 1 mL of 20 mM Tris–acetate (pH 7.5). 3. Measure absorption of 100-times diluted sample at 260 nm. 4. Calculate the concentration of the oligonucleotide in the sample using an extinction coefficient of 90  mM−1  cm−1 at 260 nm. For example, the absorption of 0.9 corresponds to (dC)12 concentration in the stock solution of 1 mM. 5. Dissolve HPLC-purified (dG)12 obtained as described in Subheading 3.1.2 in 200 mL of 0.1 M NaOH. 6. Withdraw 10 mL from the sample and add to a quartz cuvette filled with 1 mL of 20 mM Tris–acetate (pH 7.5). 7. Measure absorption of 100-times diluted sample at 260 nm. 8. Calculate the concentration of the oligonucleotide in the sample using an extinction coefficient of 120 mM−1 cm−1 at 260 nm. For example, the absorption of 0.6 corresponds to (dG)12 concentrations in the stock solution of 0.5 mM. 9. Mix proper volumes of (dG)12 and (dC)12 samples (see above) to obtain a solution having equal final concentrations of both the above oligonucleotides. The final volume of the mixture should be 200–400  mL and the concentration of (dG)12– (dC)12 should be in the range of 10–30 mM. 10. Transfer the mixture to dialysis tubing and dialyze against 1 L of 20  mM Tris–acetate (pH 7.5) for 2  h at room temperature. 11. Withdraw 100  mL from the dialyzed sample and add to a quartz cuvette filled with 0.9  mL of 20  mM Tris–acetate (pH 7.5). 12. Measure absorption of tenfold diluted sample at 260 nm. 13. Calculate the concentration of the template–primer in the sample using an extinction coefficient of 177 mM−1 cm−1 at 260 nm. For example, the absorption of 0.177 corresponds to (dG)12–(dC)12 concentration in the stock solution of 10 mM.

Synthesis of Long DNA-Based Nanowires

129

14. Transfer the (dG)12–(dC)12 sample into several 0.5 mL plastic tubes (0.1 mL per tube). 15. Freeze the samples in a dry ice/ethanol bath and store at −18°C until ready to proceed with the DNA synthesis. 3.2. Preparation of SH-(dG)12–(dC)12-SH Template–Primer

Complete separation of SH-(dG)12 and SH-(dC)12 oligonucleotides, comprising a SH-(dG)12–(dC)12-SH template–primer, from shorter or longer oligonucleotides and oligonucleotides not containing thiol-groups is required.

3.2.1. HPLC Purification SH-(dC)12

1. Transfer ~1  mg of a dry oligonucleotide powder to plastic 1.5 mL plastic tube. 2. Add 1 mL 20 mM Tris–acetate (pH 7.5). 3. Shake the sample and vortex vigorously for 2 min; incubate at room temperature for 30 min and vortex again. 4. Centrifuge the sample for 2  min at 5,000 × g at room temperature in order to get rid of insoluble compounds that might be present in the oligonucleotide preparation. 5. Transfer the entire supernatant to a new 1.5  mL plastic tube. 6. Add 25 mL of 0.4 M DTT. 7. Incubate the sample for 40 min at room temperature. 8. Connect an ion-exchange PolyWax LP column to the HPLC system. 9. Equilibrate the column with 20  mL of 2  mM Tris–acetate (pH 7.5) at a flow rate of 0.8  mL/min at room temperature. 10. Load 150 mL of the oligonucleotide sample at a flow rate of 0.8 mL/min. Do not overload the column; the large sample volume can significantly reduce the separation efficiency. 11. Elute the oligonucleotide with a linear NaCl gradient from 0 to 0.5 M for 60 min at a flow rate of 0.8 mL/min at room temperature. Monitor the elution by measuring absorbance at 260 nm. 12. Collect the fraction containing SH-(dC)12. Total volume of the fraction should be ~1.5 mL. 13. Equilibrate Sephadex G-25 DNA-Grade column with 30 mL of 2 mM Tris–acetate (pH 7.5) at room temperature. 14. Load 1.5  mL of the oligonucleotide sample obtained from the ion-exchange column (see step 12). Allow the sample to enter the column completely. Add 1.5  mL of 2  mM Tris– acetate (pH 7.5). Allow the buffer to enter the column.

130

Kotlyar

15. Place 2  mL plastic tube under the column; add 2.0  mL of Tris–acetate (pH 7.5) buffer and collect the eluant. 16. Transfer the solution into two 1.5 mL plastic tubes (1.0 mL per tube). 17. Freeze the sample in a dry ice/ethanol bath and lyophilize it to dryness. It takes approximately 15  h to completely lyophilize the sample. 18. Store the dry sample at −18°C. 3.2.2. HPLC Purification of SH-(dG)12

1. Transfer ~1  mg of a dry oligonucleotide powder to plastic 1.5 mL plastic tube. 2. Add 1 mL of 0.1 M NaOH. 3. Shake the sample and vortex vigorously for 2 min. 4. Centrifuge the sample for 2  min at 5,000 × g at room temperature in order to get rid of insoluble compounds that might be present in the oligonucleotide preparation. 5. Transfer the entire supernatant to a new 1.5  mL plastic tube. 6. Add 25 mL of 0.4 M DTT. 7. Incubate the sample for 30 min at room temperature. 8. Connect an ion-exchange HiTrap Q HP column to the HPLC system. 9. Equilibrate the column with 0.1 M NaOH at a flow rate of 0.7 mL/min at room temperature. 10. Load 150 mL of the oligonucleotide sample at a flow rate of 0.7 mL/min. 11. Elute the oligonucleotide in 0.1 M NaOH with a linear NaCl gradient from 0.5 to 1  M for 200  min at a flow rate of 0.7  mL/min at room temperature. Monitor the elution by measuring absorbance at 260 nm. 12. Collect the fraction containing SH-(dG)12. Total volume of the fraction should be ~3 mL. 13. Equilibrate Sephadex G-25 DNA-Grade column with 30 mL of 2 mM Tris–acetate (pH 7.5) at room temperature. 14. Load 3 mL of the oligonucleotide sample obtained from the ion-exchange column (see step 12). Allow the sample to enter the column completely. 15. Place 5 mL plastic tube under the column; add 3 mL of 2 mM Tris–acetate (pH 7.5) buffer and collect the eluant. 16. Transfer the solution into three 1.5 mL plastic tubes (1 mL per tube).

Synthesis of Long DNA-Based Nanowires

131

17. Freeze the sample in a dry ice/ethanol bath and lyophilize the sample to dryness. It takes approximately 15 h to completely lyophilize the sample. 18. Store the dry sample at −18°C. 3.2.3. Annealing of Purified SH-(dC)12 and SH-(dG)12

1. Dissolve HPLC-purified SH-(dC)12 obtained as described in Subheading 3.2.1 in 200 mL of 0.1 M NaOH. 2. Withdraw 10 mL from the sample and add to a quartz cuvette filled with 1 mL of 20 mM Tris–acetate (pH 7.5). 3. Measure absorption of 100-times diluted sample at 260 nm. 4. Calculate the concentration of the oligonucleotide in the sample using an extinction coefficient of 90  mM−1  cm−1 at 260 nm. For example, the absorption of 0.9 corresponds to SH-(dC)12 concentration in the stock solution of 1 mM. 5. Dissolve HPLC-purified SH-(dG)12 obtained as described in Subheading 3.2.2 in 200 mL of 0.1 M NaOH. 6. Withdraw 10 mL from the sample and add to a quartz cuvette filled with 1 mL of 20 mM Tris–acetate (pH 7.5). 7. Measure absorption of 100-times diluted sample at 260 nm. 8. Calculate the concentration of the oligonucleotide in the sample using an extinction coefficient of 120 mM−1 cm−1 at 260 nm. For example, the absorption of 0.6 corresponds to SH-(dG)12 concentration in the stock solution of 0.5 mM. 9. Mix proper volumes of SH-(dG)12 and SH-(dC)12 samples (see above) to obtain a solution having equal final concentrations of  both the above oligonucleotides. The final volume of the ­mixture should be 200–400  mL and the concentration of SH-(dG)12–(dC)12-SH should be in the range of 10–30 mM. 10. Add 2.5 mL of 0.4 M DTT per each 100 mL the mixture and incubate at room temperature for 30 min. 11. Transfer the sample (200–400 mL) to dialysis tubing and dialyze against 100 mL of 20 mM Tris–acetate (pH 7.5) buffer containing 2 mM DTT for 1 h at room temperature. Change the dialysis buffer and continue dialysis for one more hour at room temperature. 12. Withdraw 50 mL from the sample and add to a quartz cuvette filled with 1 mL of 20 mM Tris–acetate (pH 7.5). 13. Measure absorption of 20 times diluted sample at 260 nm. 14. Calculate the concentration of the template–primer in the sample using an extinction coefficient of 177 mM−1 cm−1 at 260 nm. For example, the absorption of 0.177 corresponds to SH-(dG)12–(dC)12-SH concentration in the stock solution of 20 mM.

132

Kotlyar

15. Transfer the SH-(dG)12–(dC)12-SH sample into several 0.5 mL plastic tubes (0.1 mL per tube). 16. Freeze the samples in a dry ice/ethanol bath and store at −18°C until ready to proceed with the DNA synthesis. 3.3. Preparation of Poly(dG)–Poly(dC) 3.3.1. Enzymatic Synthesis of Poly(dG)–Poly(dC)

The method described here is different from classical PCRmethods of DNA synthesis. It is based on the unique property of DNA Polymerase (Klenow exo− fragment) to extend blunt-ended poly(dG)–poly(dC) molecules in the presence of dGTP and dCTP (see Note 3). 1. Prepare the mix for the DNA synthesis. Combine the following reagents in a 0.5 mL microcentrifuge plastic tube for each reaction: 85.25 mL of deionized/filtered water, 6 mL of 1 M Phosphate buffer (pH 7.5), 0.5 mL of 1 M MgCl2, 1.5 mL of 100 mM dCTP, 1.5 mL of 100 mM dGTP and 1.25 mL of 0.4 M DTT, for a total volume of 0.1 mL (you may scale up or down accordingly). Mix well by vortexing. 2. Add 2  mL of (dG)12–(dC)12 template/primer. Mix well by vortexing. 3. Add 2 mL of Klenow exo−, mix well by vortexing and incubate the reaction at 37°C in an air dry bath for 1  h. One-hour incubation leads to synthesis of approximately 2,000 base pairs poly(dG)–poly(dC) molecules (see Fig.  3). You may change the amount of bases in the DNA accordingly by extending or reducing the incubation time. 4. Add 2 mL of 1 M EDTA to terminate the reaction and vortex the sample.

Fig. 3. Time course of poly(dG)–poly(dC) synthesis reaction. Polymerase extension assay was performed as described in Subheading 3.3.1 in the presence of 0.2 mM (dG)12–(dC)12 and 20 mg/ml of Klenow exo−; the incubation was set at 37°C. Aliquots were withdrawn each 15 min for 2 h 15 min. (a) The reaction products were resolved on 1% agarose gel and stained with ethidium bromide. The marker bands of 1 kb DNA ladder (lane 1) are indicated to the left. Time-dependent products for 15, 30, 45, 60, 75, 90, 105, 120, and 135 min of the synthesis (lanes 2–10). (b) Dependence of the polymer length (in kb) estimated from (a) on the time of synthesis.

Synthesis of Long DNA-Based Nanowires 3.3.2. HPLC Purification of Synthesized Poly(dG)–Poly(dC)

133

In order to completely separate synthesized Poly(dG)–Poly(dC) from nucleotides, the template–primer, Klenow exo−, and other reaction components of the synthesis we recommend to use sizeexclusion HPLC. 1. Connect TSK-gel G-DNA-PW HPLC column to the HPLC system (see Note 4). 2. Equilibrate the column with 20 mM Tris–acetate (pH 8.0) at a flow rate of 0.5 mL/min at room temperature. 3. Load 100  mL of poly(dG)–poly(dC) sample obtained as described above (Subheading  3.3.1) at a flow rate of 0.5 mL/min. 4. Elute the DNA in 20 mM Tris–acetate (pH 8.0) at a flow rate of 0.5 mL/min at room temperature. Monitor the elution by measuring absorbance at 260 nm. 5. Collect the DNA fraction from the column. Total volume of the fraction should be ~1 mL. 6. Withdraw 100 mL from the DNA sample and add to a quartz cuvette filled with 1 mL of 20 mM Tris–acetate (pH 8.0). 7. Measure absorption of tenfold diluted sample at 260 nm. 8. Calculate the concentration of the DNA in the sample using an extinction coefficient of 14.8 mM−1 cm−1 at 260 nm for a GC pair. For example, the absorption of 0.148 corresponds to GC concentration in the stock solution of 0.1 mM. 9. Transfer the solution into several 0.5  mL plastic tubes (0.1 mL per tube) (see Note 5). 10. Freeze the DNA samples in a dry ice/ethanol bath and store at −18°C.

3.4. Preparation of SH-Poly(dG)– Poly(dC)-SH 3.4.1. Enzymatic Synthesis of SH-Poly(dG)–Poly(dC)-SH

The method enables to obtain homogeneous population of SH-poly(dG)–poly(dC)-SH characterized by high affinity to gold surfaces and electrodes. 1. Prepare the mix for the DNA synthesis. Combine the following reagents in a 0.5 mL microcentrifuge plastic tube for each reaction: 67 mL of deionized water, 6 mL of 1 M Phosphate buffer (pH 7.5), 0.5 mL of 1 M MgCl2, 1.5 mL of 100 mM dCTP, 1.5 mL of 100 mM dGTP and 1.5 mL of 0.4 M DTT, for a total volume of 0.1  mL (you may scale up or down accordingly). Mix well by vortexing. 2. Add 20  mL of SH-(dG)12–(dC)12-SH template/primer. Mix well by vortexing. 3. Add 2 mL of Klenow exo−, mix well by vortexing and incubate the reaction at 37°C in an air dry bath for 1  h. One-hour incubation leads to synthesis of approximately 500 base pairs

134

Kotlyar

of SH-poly(dG)–poly(dC)-SH molecules. You may change the amount of bases in the DNA accordingly by extending or reducing the incubation time (see Note 6). 4. Add 2 mL of 1 M EDTA to terminate the reaction and vortex the sample. 3.4.2. HPLC Purification of Synthesized SH-Poly(dG)–Poly(dC)-SH

1. Connect TSK-gel G-DNA-PW HPLC column to the HPLC system. 2. Equilibrate the column with 20 mM Tris–acetate (pH 7.5) at a flow rate of 0.5 mL/min at room temperature. 3. Load 100 mL of SH-poly(dG)–poly(dC)-SH sample obtained as described above (Subheading  3.4.1) at a flow rate of 0.5 mL/min. 4. Elute the DNA in 20 mM Tris–acetate (pH 7.5) at a flow rate of 0.5 mL/min at room temperature. Monitor the elution by measuring absorbance at 260 nm. 5. Collect the DNA fraction from the column. Total volume of the fraction should be ~1 mL. 6. Withdraw 100 mL from the DNA sample and add to a quartz cuvette filled with 0.9 mL of 20 mM Tris–acetate (pH 7.5). 7. Measure absorption of tenfold diluted sample at 260 nm. 8. Calculate the concentration of the DNA in the sample using an extinction coefficient of 14.8 mM−1 cm−1 at 260 nm for a GC pair. For example, the absorption of 0.148 corresponds to GC concentration in the sample of 0.1 mM. 9. Transfer the solution into several 0.5  mL plastic tubes (0.1 mL per tube) (see Note 7). 10. Freeze the DNA samples in a dry ice/ethanol bath and store at −18°C.

3.5. Preparation of Poly(dG-dG)–Poly(dC)

The method of poly(dG-dG)–poly(dC) synthesis described here is based on the extension of the G-strand of the poly(dG)– poly(dC) by the Klenow exo− fragment of DNA polymerase I, under conditions when only the G-strand is allowed to grow.

3.5.1. Enzymatic Synthesis of Poly(dG-dG)–Poly(dC)

1. Prepare the mix for the DNA synthesis. Combine the following reagents in a 0.5 mL microcentrifuge plastic tube for each reaction: 74.7 mL of deionized water, 6 mL of 1 M Phosphate buffer (pH 7.5), 0.33 mL of 1 M MgCl2, 0.5 mL of 100 mM dGTP, and 1.5  mL of 0.4  M DTT, for a total volume of 0.1 mL (you may scale up or down accordingly). Mix well by vortexing. 2. Add 15  mL of 1  mM (in base pairs) poly(dG)–poly(dC) ­solution. Mix well by vortexing.

Synthesis of Long DNA-Based Nanowires

135

3. Add 2 mL of Klenow exo−, mix well by vortexing and incubate the reaction at 37°C in an air dry bath for 4 h. 4. Add 1 mL of 1 M EDTA to terminate the reaction and vortex the sample. 3.5.2. HPLC Purification of Synthesized Poly (dG-dG)–Poly(dC)

1. Connect TSK-gel G-DNA-PW HPLC column to the HPLC system. 2. Equilibrate the column with 20 mM Tris–acetate (pH 7.5) at a flow rate of 0.5 mL/min at room temperature. 3. Load 100 mL of poly(dG-dG)–poly(dC) sample obtained as described above (Subheading  3.5.1) at a flow rate of 0.5 mL/min. 4. Elute the triplex DNA in 20 mM Tris–acetate (pH 7.5) at a flow rate of 0.5 mL/min at room temperature. Monitor the elution by measuring absorbance at 260 nm. 5. Collect the DNA fraction from the column. Total volume of the fraction should be ~1 mL. 6. Withdraw 100 mL from the DNA sample and add to a quartz cuvette filled with 0.9 mL of 20 mM Tris–acetate (pH 7.5) buffer. 7. Measure absorption of tenfold diluted sample at 260 nm. 8. Calculate the concentration of the DNA in the sample using an extinction coefficient of approximately 20  M−1  cm−1 at 260 nm for a GGC triad. For example, the absorption of 0.2 corresponds to GGC concentration in the sample of 0.1 mM. 9. Transfer the solution into several 0.5  mL plastic tubes (0.1 mL per tube). 10. Freeze the DNA samples in a dry ice/ethanol bath and store at −18°C.

3.6. Preparation of G4 (Quadruple)-DNA 3.6.1. HPLC Purification of (dC)20

1. Transfer ~1  mg of a dry oligonucleotide powder to plastic 1.5 mL plastic tube. 2. Add 1 mL of deionized/filtered water. 3. Shake the sample and vortex vigorously for 2 min; incubate at room temperature for 30 min and vortex again. 4. Centrifuge the sample for 2  min at 5,000 × g at room temperature in order to get rid of insoluble compounds that might be present in the oligonucleotide preparation. 5. Transfer the entire supernatant to a new 1.5 mL plastic tube. 6. Connect an ion-exchange PolyWax LP column to the HPLC system.

136

Kotlyar

7. Equilibrate the column with 20  mL of 20  mM Phosphate buffer (pH 7.5), 10% acetonitrile, at a flow rate of 0.8 mL/min at room temperature. 8. Load 150 mL of the oligonucleotide sample at a flow rate of 0.8 mL/min. Do not overload the column; the large sample volume can significantly reduce the separation efficiency. 9. Elute the oligonucleotide with a linear gradient from 0.02 to 0.5  M Phosphate buffer (pH 7.5), 10% acetonitrile, for 60 min at a flow rate of 0.8 mL/min at room temperature. Monitor the elution by measuring absorbance at 260 nm. 10. Collect the DNA fraction from containing (dC)20. Total volume of the fraction should be ~1 mL. 11. Equilibrate Sephadex NAP-25 DNA-Grade column with 30 mL of 2 mM Tris–acetate (pH 7.5) at room temperature. 12. Load 1 mL of the oligonucleotide sample obtained from the ion-exchange column (see step 10). Allow the sample to enter the column completely. Add 2 mL of 2 mM Tris–acetate (pH 7.5). Allow the buffer to enter the column. 13. Place 2 mL plastic tube under the column; add 2 mL of 2 mM Tris–acetate (pH 7.5) buffer and collect the eluant. 14. Transfer the solution into four 1.5 mL plastic tubes (0.5 mL per tube). 15. Freeze the sample in a dry ice/ethanol bath and lyophilize the sample to dryness. It takes approximately 15 h to completely lyophilize the sample. 16. Store the dry sample at −18°C. 3.6.2. Enzymatic Synthesis of Poly(dG)–n(dC)20

Klenow exo− fragment of DNA Polymerase is capable of producing long double-stranded poly(dG)–n(dC)20 molecules composed of a long continuous dG-strand and relatively short dC-­ oligonucleotides not covalently connected to each other in the presence of dGTP and (dC)20. 1. Prepare the mix for the DNA synthesis. Combine the ­following reagents in a 0.5  mL microcentrifuge plastic tube for each ­reaction: 78.5 mL of deionized water, 6 mL of 1 M Phosphate buffer (pH 7.5), 0.5 mL of 1 M MgCl2, 1.5 mL of 100 mM dGTP and 1.5 mL of 0.4 M DTT, for a total volume of 0.1 mL (you may scale up or down accordingly). Mix well by vortexing. 2. Dissolve HPLC-purified (dC)20 obtained as described in Subheading  3.6.1 in 200  mL of 20  mM Phosphate buffer (pH 7.5). 3. Withdraw 1 mL from the sample and add to a quartz cuvette filled with 1 mL of 20 mM Phosphate buffer (pH 7.5).

Synthesis of Long DNA-Based Nanowires

137

4. Measure absorption of 1,000-times diluted sample at 260 nm. 5. Calculate the concentration of the oligonucleotide in the sample using an extinction coefficient of 144 mM−1 cm−1 at 260 nm. For example, the absorption of 0.144 corresponds to (dC)20 concentration in the sample of 1 mM. 6. Add 10  mL of 1  mM (dC)20 to a final concentration of 100 mM. 7. Add 2  mL of (dG)12–(dC)12 template/primer. Mix well by pipetting. 8. Add 2 mL of Klenow exo−, mix well by vortexing and incubate the reaction at 37°C in an air dry bath for 1  h. Two-hour incubation leads to synthesis of poly(dG)–n(dC)20 molecules composed of approximately 2,000 base-long G-strand. You may change the amount of bases in the DNA accordingly by extending or reducing the incubation time. 9. Add 2 mL of 1 M EDTA to terminate the reaction and vortex the sample. 3.6.3. HPLC Purification of Synthesized Poly(dG)–n(dC)20

1. Connect TSK-gel DNA-G-DNA PW HPLC column to the HPLC system. 2. Equilibrate the column with 20 mM Tris–acetate (pH 7.5) at a flow rate of 0.5 mL/min at room temperature. 3. Load 100  mL of poly(dG)–n(dC)20 sample obtained as described above (Subheading  3.6.2) at a flow rate of 0.5 mL/min. 4. Elute the DNA in 20 mM Tris–acetate (pH 7.5) at a flow rate of 0.5 mL/min at room temperature. Monitor the elution by measuring absorbance at 260 nm. 5. Collect the DNA fraction from the column. Total volume the fraction should be ~1 mL.

3.6.4. Preparation of Poly(dG)

At pH higher than 12.5 the poly(dG)- and the (dC)20-fragments composing poly(dG)–n(dC)20 are separated from each other and are eluted separately from the HPLC column. 1. Connect TSK-gel G-DNA-PW HPLC column to the HPLC system. 2. Equilibrate the column with 0.1 M NaOH solution at a flow rate of 0.5 mL/min at room temperature. 3. Transfer 100  mL of poly(dG)–n(dC)20 sample obtained as described above (Subheading 3.6.3) to a 0.5 mL microcentrifuge plastic tube. 4. Add 15 mL of 1 M NaOH and incubate for 10 min at room temperature.

138

Kotlyar

5. Load the sample onto the column at a flow rate of 0.5 mL/min. 6. Elute the DNA in 0.1 M NaOH at a flow rate of 0.5 mL/min at room temperature. Monitor the elution by measuring absorbance at 260 nm. 7. Collect a poly(dG) fraction eluted between 14 and 16 min. Total volume of the fraction should be ~1 mL. 3.6.5. Preparation of G4-DNA

Folding of G-strands into G4-structures is taking place spontaneously upon pH reduction during chromatography of the alkaline strands solution on a Sephadex G-25 column. 1. Equilibrate a Sephadex G-25 column with 5  mL of 2  mM Tris–acetate (pH 8.0) at room temperature. 2. Load 0.5 mL of the alkaline G-strand sample obtained from the TSK-gel G-DNA-PW HPLC column (see Sub­ heading  3.6.4). Allow the sample to enter the column ­completely. Add 0.2  mL of 2  mM Tris–acetate (pH 8.0). Allow the buffer to enter the column. 3. Place 1.5 mL plastic tube under the column; add 0.7 mL of 2 mM Tris–acetate (pH 8.0) buffer and collect the eluant. 4. Measure absorption of the sample at 260 nm. 5. Calculate the concentration of the DNA (in tetrads) in the sample using an extinction coefficient of 36  mM−1  cm−1 at 260 nm. For example, the absorption of 0.36 corresponds to 10 mM G4-DNA. 6. The sample can be stored for 2–3 days at 4°C. Longer storage is not recommended. Do not freeze the sample.

4. Notes 1. Complete purification of (dG)12, (dC)12, SH-(dG)12, SH-(dC)12 comprising a (dG)12–(dC)12 and SH-(dG)12– (dC)12-SH ­template–primers from shorter or longer oligonucleotides that are usually present in minor quantities in commercial preparations is required. If primed by nonpurified template–primers, the synthesis yields DNA molecules with large length variability. 2. Steps 8–13 in Subheadings 3.1.1 and 3.1.2 can be repeated several times in order to obtain larger quantities of purified oligonucleotides for a large-scale synthesis of the poly(dG)–poly(dC).

Synthesis of Long DNA-Based Nanowires

139

3. The protocol of poly(dG)–poly(dC) synthesis (see Subheading  3.3) can be adapted for synthesis of poly(dA)– poly(dT), a double-stranded polymer composed of poly(dA)and poly (dT)-homopolymer strands of equal length. 4. If the length of the synthesized poly(dG)–poly(dC) is shorter than 1  Kbp, use TSK-gel G-5000-PW HPLC column (7.8 × 300  mm, TosoHaas, Japan) instead of TSK-gel G-DNA-PW HPLC column (7.8 × 300  mm, TosoHaas, Japan) for the HPLC purification of the DNA. 5. The G-containing structures are unstable at low pH and undergo acid hydrolysis. We thus recommend storage of the DNA samples at pH 8–8.5 at 4°C. 6. The rate of SH-poly(dG)–poly(dC)-SH synthesis is slower than that of non-thiolated poly(dG)–poly(dC) polymer. Relatively large quantities of the enzyme should therefore be added into the assay in order to obtain long SH-poly(dG)– poly(dC)-SH molecules. 7. Thiol-groups can undergo spontaneous oxidation to disulfides. We recommend to store SH-poly(dG)–poly(dC)-SH in the presence of 1  mM DTT to avoid disulfides formation. The molecules should be separated from DTT prior to deposition on gold surfaces. This can be done by passing the DNA sample through Sephadex G-25 DNA-Grade column equilibrated with 2 mM Tris–acetate (pH 7.5).

Acknowledgments This work was supported by the EC through the contracts IST2001-38951 (“DNA-Based Nanowires”) and FP6-029192 (“DNA-Based Nanodevices”). References 1. Porath, D., Bezryadin, A., de Vries, S. and Dekker, C. (2000) Direct measurement of electrical transport through DNA molecules. Nature, 403, 635–638. 2. Hwang, J. S. K., Kong, J., Ahn, D. G., Lee, S., Ahn, D. J. S. and Hwang, W. (2002) Electrical transport through 60 base pairs of poly(dG)-poly(dC) DNA molecules. Appl. Phys. Lett., 81, 1134–1136. 3. Hennig, D., Starikov ,E. B., Archilla, J. F. R. and Palmero, F. (2004) Charge transport in poly(dG)-poly(dC) and poly(dA) poly(dT) DNA polymers. J. Biol. Phys., 30, 227–238.

4. Yi, J. (2003) Conduction of DNA molecules: A charge-ladder model. Physic. Rev. B. 68, 193103. 5. Lee, H.-Y., Tanaka, H., Otsuka, Y., Yoo, K.-H., Lee, J.-O. and Kawai,T. (2002) Control of electrical conduction in DNA using oxygen hole doping. App. Phys. Lett., 80, 1670–1672. 6. Yoo, K.-H., Ha, D.H., Lee, J.-O., Park, J.W., Kim, J., Kim, J.J., Lee, H.-Y., Kawai, T. and Choi, H.-Y. (2001) Electrical conduction through Poly(dA)–Poly(dT) and Poly(dG)– Poly(dC)DNAmolecules. Phys. Rev. Lett., 87, 198102.

140

Kotlyar

7. Kotlyar, A. B., Borovok, N., Molotsky T., Fadeev L., and Gozin M. (2005) In Vitro synthesis of uniform Poly(dG)-Poly(dC) by Klenow exo– fragment of Polymerase I. Nucl. Acid Res. 33, 525–535. 8. Nuzzo, R. G. and Allara, D. L. (1983) Adsorption of bifunctional organic disulfides on gold surfaces. J. Am. Chem. Soc., 105, 4481–4483. 9. Sellers, H., Ulman, A., Shnidman, Y. and Eilerss, J.E. (1993) Structure and binding of alkanethiolates on gold and silver surfaces: implications for self-assembled monolayers. J. Am. Chem. Soc., 115, 9389–9401. 10. Hegner, M., Wagner, P. and Semenza, G. (1993) Immobilizing DNA on gold via thiol modification for atomic force microscopy imaging in buffer solutions. FEBS Lett., 336, 452–456. 11. Frank-Kamenetskii, M. D. and Mirkin, S. M. (1995) Triplex DNA structures. Annu. Rev. Biochem., 64, 65–95. 12. Sun, J. S., Garestier, T. and Helene, C. (1996) Oligonucleotide directed triple helix formation. Curr. Opin. Struct. Biol., 6, 327–333. 13. Radhakrishnan, I. and Patel, D. J. (1994) DNA triplexes: solution structures, hydration sites, energetics, interactions, and function. Biochemistry, 33, 11405–11416. 14. Kotlyar, A. B., Borovok, N., Molotsky, T, Klinov, D., Dwir, B. and Kapon E. Synthesis of novel poly(dG)-poly(dG)-poly(dC) triplex structure by Klenow exo- fragment of DNA polymerase I. 2005 Nucl. Acid Res. 33, 6515–6521. 15. Kerwin, S. M. (2000) G-Quadruplex DNA as a target for drug design. Curr. Pharmaceutic. Design, 6, 441–478.

16. Davis, J. T. (2004) G-quartets 40 years later: From 50-GMP to molecular biology and supramolecular chemistry, Angew. Chem. Intl. Ed. 43, 668–698. 17. Keniry M. A. (2001) Quadruplex Structures in Nucleic Acids. Biopolymers, 56, 123–146. 18. Parkinson, G. N., Lee, M. P. and Neidle, S. (2002) Crystal structure of parallel quadruplexes from human telomeric DNA. Nature, 417, 876–880. 19. Burge, S., Parkinson, G. N., Hazel, P., Todd, A. K. and Neidle, S. (2006) Quadruplex DNA: sequence, topology and structure. Nucleic Acids Res., 34, 5402–5415. 20. Sen, D. and Gilbert, W. (1992) Novel DNA superstructures formed by telomere-like oligomers. Biochemistry, 31, 65–70. 21. Marsh, T. C., Vesenka, J. and Henderson, E. (1995) A new DNA nanostructure the G-wire imaged by scanning probe microscopy. Nucleic Acids Res., 23, 696–700. 22. Kotlyar, A. B., Borovok, N., Molotsky, T., Cohen, H., Shapir, E. and Porath, D. (2005) Long monomolecular guanine-based nanowires, Adv. Mater. 17, 1901–1905. 23. Borovok, N, Molotsky, T, Ghabboun, J, Porath, D. and Kotlyar, A. (2008) Efficient procedure of preparation and properties of long uniform G4-DNA nanowires. Anal. Biochem. 374, 71–78. 24. Cohen, H., Sapir, T., Borovok, N., Molotsky, T., Di Felice, R., Kotlyar, A. B. and Porath, D. (2007) Polarizability of G4-DNA observed by electrostatic force microscopy measurements. Nano Letters, 7, 981–986.

Chapter 10 G-Wire Synthesis and Modification with Gold Nanoparticle Christian Leiterer, Andrea Csaki, and Wolfgang Fritzsche Abstract DNA molecules are well known for containing the genetic information of an individual. Furthermore, DNA is a biopolymer with the potential of building up nanoscale structures. These structures can be addressed sequence specifically and, therefore, they allow connecting and arranging with subnanometer accuracy. The extended work of the group of Nadrian Seeman (Nature 421:427–431, 2003) has shown that the self-assembly of DNA molecules offers great potential for the creation of bottom-up nanostructures for nanoelectronics, biosensors, and programmable molecular machines. Rothemund (Nature 440:297– 302, 2006) has shown that it is possible to generate a wide variety of 2D nanostructures by the assembly of synthetic desoxyoligonucleotides and M13mp18 DNA via Watson–Crick base pairing. Furthermore, DNA can form three- and four-stranded structures which offer even more possibilities for molecular construction. This chapter will deal with four-stranded DNA structures (G-wires) created from 10-bp deoxynucleotide units. Our focus will be especially on the synthesis, individualization, modification with gold nanoparticles, and characterization by high-resolution scanning force microscopy (AFM). Key words: G-wire, Quadruplex, Gold nanoparticle, AFM, Nanowires

1. Introduction G-wires assemble thanks to DNA–DNA hybridization of four guanine (G)-nucleotides (3, 4). In nature, such G-quartet motifs can be found at the telomeres of chromosomes and in ribozymes where they provide the DNA with extra structural stability against enzymatic digestion and chemical reactions. Artificial structures based on this motif can be prepared either by intermolecular (5) or intramolecular (6) G-quartet formation using oligonucleotides containing predominantly (or solely) guanines. Hydration layer scanning tunnel microscopy (HLSTM) (7) imaging has suggested that G-wires might function as semiconductors (8) due to base stacking of the G-quartets and the caged monovalent cations. This potential for conduction

Giampaolo Zuccheri and Bruno Samorì (eds.), DNA Nanotechnology: Methods and Protocols, Methods in Molecular Biology, vol. 749, DOI 10.1007/978-1-61779-142-0_10, © Springer Science+Business Media, LLC 2011

141

142

Leiterer, Csaki, and Fritzsche

and the formation of long stable structures makes G-wire interesting not only for molecular construction, but also for nanoelectronic applications (9, 10). The oligonucleotide with the sequence 5¢-GGGGTTGGGG-3¢ was used for the assembling of G-wires in this protocol. The four guanines on both ends can hybridize with three other oligonucleotides each, resulting in a polymerization reaction that leads to long (several tens up to hundreds of nanometers) G-rich quadruplex structures (5). The polymerization reaction is controllable by hybridization temperature, time, and buffer composition (11). Using biotinylated oligonucleotides in combination with streptavidin-conjugated gold nanoparticles or positively char­ ged gold, nanoparticles were specifically attached either to the ends or unspecifically to the backbone of the G-wire (12).

2. Materials 2.1. Synthesis and Immobilization of G-Wire

1. 10-nt oligonucleotide with the sequence 5¢-GGGGTTGGGG-3¢, (G4), from Jena Bioscience GmbH (Jena, Germany). 2. Synthesis buffer (P1): 50  mM NaCl, 50  mM  Tris–HCl, pH 7.4, and 10 mM MgCl2 (see Note 1). 3. Dilution buffer (P2): 10 mM Tris–HCl, pH 7.4, and 1 mM MgCl2. 4. Mica sheets for AFM, Hi-Grade quality (Plano Planet GmbH, Wetzlar, Germany). 5. Photolithographically structured (13–15) microelectrode chips on thermally oxidized silicon (1  mm). A 3–5-nm titanium adhesion layer and a 200-nm gold layer were deposited by sputtering, followed by a lift-off process. 6. Chip activation solution: 1:1 ratio mix of 30% H2O2 (puriss p.a.) and 25% NH4OH (puriss p.a.).

2.2. Isolation and Growth of G-Wire

1. G-buffer: 50 mM KCl, 10 mM MgCl2, and 50 mM Tris–HCl, pH 7.4.

2.3. Surface Passivation and Gold Nanoparticle Modification of G-Wires Using GENOgold

1. Mica sheets for AFM, Hi-Grade quality (Plano Planet GmbH, Wetzlar, Germany). 2. 1× PBS: 137  mM NaCl, 10  mM phosphate, and 2.7  mM KCl, pH 7.4. 3. PEG solution: 1 M PEG. 4. MgCl2 solution: 5 mM MgCl2. 5. Mg–acetate solution: 1 M Mg(CH3COO)2. 6. Genogold (BBI, UK): 6°× 10 11  particles/ml.

17–23  nm

in

diameter,

G-Wire Synthesis and Modification with Gold Nanoparticle

2.4. Modification of Biotinylated G-Wire with Streptavidin-Modified Gold Nanoparticle

143

1. Biotinylated oligonucleotide [G4T2G4-biotin, Jena Bioscience GmbH (Jena, Germany)]. 2. Streptavidin-conjugated 1.6°× 1014 particles/ml).

gold

nanoparticles

(5 

nm,

3. Methods A first objective was to synthesize long and separated G-wires, in order to immobilize them between two electrodes for conductivity measurements or further DNA construction experiments. Previous work had shown that in most cases, G-wire could only be assembled in tight network or very small single wires. The next step would be to synthesize long single G-wire for further investigations. To achieve this goal, G-wires were immobilized on different surfaces in a G-wire compatible buffer. Furthermore, G-wires were reassembled after synthesis in a repeated heating/ cooling cycle in order to get longer single G-wire structures. A second objective was the specific binding of gold nanoparticles to G-wires for further optical and electrical experiments. To this end, two strategies were used. One was direct binding of gold nanoparticles to the backbone of the G-wires using the affinity of positively charged gold nanoparticles to the negatively charged DNA backbone. Alternatively, streptavidin-conjugated gold nanoparticles were bound to G4T2G4-biotin-modified oligonucleotides which were previously integrated in the G-wire structures. 3.1. Synthesis and Immobilization of G-Wire

1. Prior to the synthesis of the G-wire, the lyophilized oligonucleotide was resuspended in ddH2O (250 mg/ml) and heated to 95°C for preventing unspecific binding between the oligonucleotides. 2. Afterward, the oligonucleotides were diluted in synthesis buffer (P1) (1:2) and incubated at 37°C for 4 days to assemble the G-wire structures. The dilution was covered with mineral oil to avoid evaporation and to keep a constant concentration during the whole assembly process. 3. After assembling, the G-wires were diluted in dilution buffer (P2) (1:20, final oligo concentration 6.25 ng/ml) and stored for up to 2 days at room temperature for immediate use, or at −20°C for storage. 4. For the immobilization of G-wires on mica sheets, the mica surface was activated by cleavage just before spreading (see Note 2).

144

Leiterer, Csaki, and Fritzsche

5. For immobilization on silicon (thermally oxidized), the surface was activated either by incubation in chip activation solution for 1  h at 70°C or by oxygen plasma etching at 50 W for 12 min. 6. On both surfaces (mica and oxidized silicon), the G-wire dilution was incubated for at least 4 min to achieve binding to the surface (see Note 3). Afterward, the mica sample was washed with 1 ml of ddH2O in order to remove remaining salt from the surface (see Note 4). 7. The G-wire can be visualized by AFM imaging in air, using a Nanoscope III with a Dimension 3100 AFM head. All images were taken in tapping mode™. In order to get more stable images by minimizing surface charges, the samples were stored at up to 1 week (see Note 5, Fig. 1). 3.2. Isolation and Growth of G-Wire

1. For the isolation and growth of the synthesized G-wire, the salt concentration was raised from synthesis buffer level to the G-buffer level for extra stabilization of the G-wire (see Note 6). 2. The G-wire suspension was diluted 1:2 and 1:4 (final oligo concentration 3 and 1.5 ng/ml, respectively) in order to get single wires. 3. For the growth of the G-wire, the suspension was heated and cooled (95°C/55°C) for seven cycles in order to get long G-wire structures (see Note 7). 4. For immobilization on mica, the mica surface is activated by cleavage just prior to spreading. 5. The G-wire dilution was incubated for at least 4  min to achieve the binding to the mica surface. 6. The samples were stored at room temperature for a few days in order to get dry and non-charged surface. Visualization of the G-wire was performed by AFM imaging in air, using a Nanoscope III with a Dimension 3100 AFM head. All images were taken in tapping mode™. In order to get more stable images by minimizing surface charges, the samples were stored up to 1 week (see Fig. 2).

3.3. Surface Passivation and Gold Nanoparticle Modification of G-Wires Using Genogold

1. To enable specific binding of gold nanoparticle to the G-wire, the surface must be passivated in order to prevent unspecific nanoparticle binding to the surface. 2. The mica surface was activated by cleavage just before use. 3. Then the surface was incubated in four different buffers: 1× PBS, PEG solution, MgCl2 solution, and Mg–acetate solution to saturate the negative charges on the surface. The surface was then washed with ddH2O and dried with gaseous N2.

G-Wire Synthesis and Modification with Gold Nanoparticle

145

Fig. 1. Immobilized G-wire on different surfaces and different conditions visualized by AFM imaging (all in P1/P2 mixture). (a) Negative control: AFM micrograph of the (G4T2G4) oligonucleotide spread on mica before G-wire synthesis; the oligonucleotide was stored at −20°C and 6.25 ng/ml. Only very short G-wire structures are evident. (b) AFM micrograph of the (G4T2G4) oligonucleotide spread on mica before G-wire synthesis; the oligonucleotide was stored at −20°C and 25 ng/ml. A carpet of G-wire formed on the mica surface. (c) Synthesized G-wires (made from 6.25 ng/ml oligonucleotide) immobilized on mica; these were forming a very tight arrangement of individual G-wire structures with few interconnections. (d) Same as (c) but immobilized on thermally oxidized silicon; such conditions lead to a significantly reduced number of immobilized G-wires compared to same conditions on mica.

4. The G-wire dilution was incubated on the mica surface for 4  min at room temperature and washed with ddH2O afterward. 5. Genogold was incubated for 15 min at room temperature on the surface and subsequently washed with ddH2O and dried with gaseous N2 (see Note 8).

146

Leiterer, Csaki, and Fritzsche

Fig. 2. Effect of heating cycle on the growth of the G-wires. G-wires were immobilized on mica visualized by AFM imaging. Changes in buffer concentration have been employed in order to get longer individual G-wire structures. (a) 4 days, 37°C synthesis at standard concentration (6.25 ng/ml) in P1/P2, single wires and network structures were formed. (b) Enlarging of G-wires synthesized for 4 days (cf. Fig. 2a) in P1/P2, using a heating/cooling cycle. Long network-like structures were formed probably due to re-hybridization of unstable loop endings. (c) 4 days, 37°C synthesized G-wires (6.25 ng/ml) in G-buffer were forming a tight network with hardly any visible end. Probably high salt concentration leads to stronger G-wire hybridization. (d) G-wires (3 ng/ml) were synthesized for 4 days at 37°C (same Fig. 2c) in G-buffer. Now the network-like structures visible in Fig. 2c are destroyed, and short single G-wires become visible. (e) G-wires (1.5 ng/ml) were synthesized for 4 days at 37°C in G-buffer, resulting in very short single G-wires at the surface.

6. Visualization of the nanoparticle-modified G-wire was realized by AFM imaging in air after a few days when all surface charges were neutralized (see Fig. 3). 3.4. Modification of Biotinylated G-Wire with Streptavidin-Modified Gold Nanoparticle

1. Biotinylated oligonucleotides (G4T2G4-biotin) were added to the previously synthesized G-wire at a concentration ranging between 10 and 100  mM (to achieve different degrees of modification) and stored at 37°C to insert a biotin modification in the G-wire. 2. For the modification of the biotinylated G-wire with streptavidin-conjugated gold nanoparticle, the dilution was incubated at room temperature for a few minutes. 3. To immobilize the nanoparticle-modified G-wire on mica, the mica surface was activated by cleavage.

G-Wire Synthesis and Modification with Gold Nanoparticle

147

Fig. 3. Binding of Genogold to G-wires immobilized on mica. Different buffers were used for the saturation of surface charges in order to passivate the surface against unspecific binding of gold nanoparticle. (a) 1× PBS-passivated surface with no G-wire visible. Gold nanoparticles were immobilized to the surface. (b) 1 M PEG-passivated surface, very short G-wires were immobilized. Gold nanoparticles were immobilized to the surface due to unspecific binding. (c) 5  mM MgCl2passivated surface. Short G-wire could be immobilized to the surface. Very few gold nanoparticles were bound unspecifically to the surface. Nearly no gold nanoparticle was bound to the G-wire. (d) 1 M Mg(CH3COO)2-passivated surface, G-wire network could be immobilized on the surface. Gold nanoparticles were incorporated in the G-wire network. Black arrows point to G-wires, white arrows point to gold nanoparticles.

4. The G-wire dilution was incubated for 4  min to provide binding to the mica surface. 5. The prepared samples were stored for drying and discharging up to 1 week. The samples were characterized by AFM imaging in air, using a Nanoscope III with a Dimension 3100 AFM head. All images were taken in tapping mode™. In order to get more stable images by minimizing surface charges, the samples were stored up to 1 week (see Fig. 4).

148

Leiterer, Csaki, and Fritzsche

Fig. 4. Streptavidin–Au-modified biotinylated G-wire on mica. G-wires were synthesized from a mixture of non-biotinylated (G4T2G4, 120 mM) and biotinylated oligonucleotides (G4T2G4-biotin) in different concentrations (10–100 mM). Afterward, streptavidin-conjugated gold nanoparticles were bound to the biotin modification on the G-wire. (a) 10 mM biotinylated oligonucleotides used for the synthesis of G-wire. The immobilized G-wire structures forming a tight network due to the small fraction (1/12) of biotinylated oligonucleotides which are causing a break in the polymerization reaction. A few streptavidin–Au nanoparticles were incorporated, (b) 20 mM biotinylated oligonucleotides, corresponding to 1/6 of the overall oligonucleotide concentration, forming a wide network of G-wires. More streptavidin–Au nanoparticles get incorporated due to the higher fraction of biotinylated oligonucleotides. (c) 50 mM biotinylated oligonucleotides (corresponding to 1/3) causing the beginning of dissociation of the network and the formation of big streptavidin–Au clusters in the G-wire network. (d) 100 mm biotinylated oligonucleotides, corresponding to a mixture of equal amounts of biotinylated and non-biotinylated oligonucleotides, do not form long G-wire structures. The biotin ending of the oligonucleotide probably hinders the polymerization reaction. Streptavidin–Au nanoparticles are evenly spread on the surface. Black arrows point to G-wires, white arrows point to gold nanoparticles.

4. Notes 1. All solutions were prepared in deionized water (ddH2O with a conductivity less than 20  mS/cm, no special filtering was necessary). 2. Cleavage was done by removing the upper mica layers from the mica sheets with a sticky tape. The result is a homogenous negatively charged and very planar surface.

G-Wire Synthesis and Modification with Gold Nanoparticle

149

3. The surface binding of G-wire is arranged by bivalent cations such as Mg2+. 4. Remaining salt from the buffer solution results in the growth of crystals on the surface, which hampers the AFM imaging; therefore, the crystals have to be removed. 5. Due to the previous surface activation, the surface remains charged even after immobilization. These remaining charges destabilize AFM imaging. It has been shown that the easiest way to get rid of the surface charges is storing the samples up to 1 week at room temperature and medium relative humidity (30% 99.5% (200 proof). 3. Acetic acid, glacial ACS reagent grade. 4. Ethanol, ACS reagent grade. 5. Acetone, ACS reagent grade. 6. Gibco™ Dulbecco’s phosphate buffer saline (PBS) 1× (no magnesium, no calcium, 2.7 mM potassium chloride, 0.14 M sodium chloride, 1.5 mM potassium phosphate, and 8 mM sodium phosphate, pH 7.4), Invitrogen; store at room temperature (RT). 7. ThermoScientific BupH™ PBS (TPBS) (0.1 M sodium phosphate and 0.15 M sodium chloride, pH 7.2); store at RT. 8. Deionized (DI) Millipore water (resistivity of 18 MWcm). 9. HS–(CH2)11–(C2H6O2)3–OH and HS–(CH2)11–(C2H6O2)3– biotin (ProChimia); store at −20°C. 10. Polyethylene glycol (PEG)-silane, 5,000 MW (mPEG5000) (LaysanBio); store at −20°C. 11. Glass syringes, metal needles for the anhydrous solvents (Popper). 12. Teflon mini-rack (Invitrogen). 13. 6-Well plates (Falcon). 14. Parafilm. 15. Streptavidin (Invitrogen); store at −20°C. 16. Chicken egg albumin (Sigma); store at 4°C. 17. Oligonucleotides (IDT): one 20-mer with a biotin functional group at the 5¢ position (5¢-/52-Bio/GTC ACT TCA GCT GAG ACG CA-3¢) and the complementary strand with a Cy3 fluorophore at the 5¢-end (5¢-/5Cy3/TGC GTC TCA GCT GAA GTG AC-3¢); store at 4°C wrapped in aluminum foil.

172

Palma et al.

2.3. Microscopy

In our laboratory, epifluorescence microscopy was performed on an inverted microscope, Olympus IX81 (Olympus) equipped with a Cascade II, 512 × 512 pixel CCD camera (Photometrics). 1. Cloning ring (Sigma/Aldrich). 2. High vacuum grease (Dow Corning).

3. Methods Throughout the steps of the method described here, a great deal of attention should be focused on the cleanliness of both the laboratory working environment and the materials and tools employed. Because of the nanometer scale of the features fabricated (and functionalized), we recommend carrying out the fabrication steps (Subheading 3.1) in an ultraclean environment, such as a clean room. Furthermore, in order to obtain a successful bio-functionalization (Subheading 3.2) of the fabricated nanopatterns, it is important to carry out the procedure rapidly with as short a time lapse as possible between steps. Moreover, all glassware and tweezers used must be dry, preferably stored in an oven at approximately 70°C, and cooled in air prior to use. It should be noted that in order to verify the validity of our approach (i.e., the controlled DNA functionalization of nanopatterned surfaces), we have used a biotinylated and Cy3-labeled dsDNA. We have hybridized in solution, prior to the attachment to the surface, a biotinylated oligonucleotide (20-mer chain) and its complementary oligonucleotide labeled with a Cy3 fluorophore. (In Subheading 4, we report the alternative procedure to immobilize biotinylated single-stranded DNA). 3.1. Metal on Glass Nanopattern Fabrication 3.1.1. Preparation and Cleaning

Cleaning is absolutely critical to the success of the procedure. Any contamination, even nanoscopic, not removed prior to patterning will result in defects in the surface passivation (see Note 1). 1. Prepare piranha solution (1/3 volume of H2O2 plus 2/3 of H2SO4). 2. Starting with a new glass coverslip, sonicate in ethanol for 2 min. 3. Blow dry with a stream of inert gas (Ar or N2) (see Note 2). 4. Dilute 7× detergent 1:4 with deionized (DI) water and bring to boiling temperature on a hot plate at 200°C (goes from cloudy to clear). 5. Immerse the slide for 2 min in boiling solution and rinse with DI water for 10 min. 6. Immerse in piranha solution and let soak for 5 min.

Controlled Confinement of DNA at the Nanoscale

173

7. Take the slide out of the piranha solution and rinse it for 10 min with DI water. 8. Rinse with ethanol. 9. Blow dry with a stream of inert gas (Ar or N2) (see Note 3). It is now possible to proceed to resist deposition. 3.1.2. Resist Deposition

Again, cleanliness is paramount here. All work should be performed in a clean room, class 10,000 or better. A bilayer of higher MW resist is spun on top of a lower molecular weight resist to aid with the subsequent metal deposition and liftoff (discussed later). The high MW top layer has a slightly different dose curve and will develop with a narrower opening, creating an overhang, which ensures proper liftoff (see Note 4). 1. Preheat a hot plate to 180°C and place cleaned samples on the hot plate for 1  min to force off any adsorbed water molecules. 2. Let cool for 10 s before placing the sample on a resist spinner chuck (see Note 5). 3. Spin lower MW resist (25K) first at 4,000 rpm for 45 s, use a ramp rate of 1,000 rpm/s (see Note 6). 4. Bake for 5 min at 180°C on a hot plate. 5. Let cool for 10 s before placing the sample on the resist spinner chuck. 6. Spin higher molecular weight resist (495K) as top layer at 4,000 rpm for 45 s, use a ramp rate of 1,000 rpm/s. 7. Bake for 5 min at 180°C. 8. Let cool for 10 s before placing on the resist spinner chuck. 9. Spin on the Aquasave conductive discharge layer at 1,000 rpm for 45 s at a ramp rate of 300 rpm/s. It is important to make sure to dab any droplets of Aquasave at the edges (particularly corners) so that the sample is completely dry before being placed in an electron-beam writer. It is now possible to proceed to the e-beam writing step.

3.1.3. Electron-Beam Pattern Writing

An electron-beam writing system is an SEM that controls beam shuttering and position to generate patterns from CAD files. Process testing will be necessary to determine optimal doses for generating the desired features (see Note 7). We use a pattern which is 50 mm by 50 mm, and consists of 1 mm register squares spaced every 10  mm, with sub-50-nm dots filling every 2  mm between them. This ensures that each individual dot is optically resolvable and discrete once functionalized and imaged with a fluorescence microscopy (see Note 8).

174

Palma et al.

1. Load samples in the e-beam writer, making sure that a good top contact is made with the mounting clips. 2. Pump down chamber. When pressure has reached or is below the specified value (in our case, 5 × 10−6 Torr), turn the beam on; use an acceleration voltage of 30 kV. 3. Check the beam currents of the spot sizes to be written within the Faraday cup. It is necessary to check beam currents each time in the Faraday cup, as currents drift over time. 4. Update the run files with the most recent beam currents to ensure that the doses are consistent. 5. Check gun tilt and stigmation, and adjust these until both are minimized (see Notes 9–11). 6. Move to the sample. Four-point focus (see Note 11) must be taken to correct for sample tilt. No sample is perfectly level, resulting in loss of focus over the writing area. Focus on the surface of the sample at four points around the region to be written, registering the points with the control software. A plane is then fit to these four points, and the focal depth interpolated and adjusted as the pattern is written (see Notes 10 and 11). 7. With all the steps taken to ensure proper focus and minimal stigmation over the area of the pattern, move to a position inside the four-point focus region. 8. Engage automated stage and beam control, and execute pattern writing in software. 9. After writing, it is necessary to remove the discharge layer, Aquasave. Rinse with DI water until all Aquasave is visibly gone. 10. Then blow dry with a stream of inert gas (Ar or N2). The samples are now ready for development. 3.1.4. PMMA Development

A cold ultrasonic development process is used to achieve nanometerscale features. The cold development sharpens resist contrast, resulting in the smallest possible features from the exposed regions. 1. Place rinsed and dry sample in a solution of 1:3 H2O:IPA at 4°C. Sonicate for 1 min in a water bath sonicator at 4°C (see Notes 12 and 13). 2. Quickly remove the sample and immerse in 100% IPA at room temperature to halt development (see Note 14). 3. Blow dry with a stream of inert gas (Ar or N2). 4. Examine pattern: large alignment marks should be visible; check further in an optical microscope at magnifications of 25–100×. The micron-sized features should be visible. It is now possible to proceed to the metal deposition step.

Controlled Confinement of DNA at the Nanoscale 3.1.5. Metal Deposition

175

In our laboratory, metal deposition is performed in a Semicore electron beam evaporator. A focused beam of electrons is used to heat a target material held in a crucible. As opposed to a thermal evaporator, the heating is highly localized to the surface and allows for precise control of the thickness (at the Ångström level). The sample is held above the target some distance away, allowing for a highly directional and uniform material flux onto the surface. 1. Place the developed samples on the sample holder in the evaporator. 2. Pump the system to less than the suggested threshold pressure and begin deposition procedure. 3. Evaporate a 1-nm adhesion layer of titanium. This is necessary for the adhesion of the Au/Pd to the glass. 4. Next, deposit 3 nm of Au/Pd on top of the titanium. 5. Follow proper power-down and venting procedures, and remove the sample. The metalized sample now consists of a layer of metal sitting atop the unexposed PMMA, with openings in the resist where the sample was exposed to electrons during the e-beam writing session (in these holes, the Ti adhesion layer and Au/Pd are deposited on the glass surface): see Fig. 1.

Fig. 1. Scanning electron microscope (JEOL JSM-5600 LV) image of nanopattern holes prior to liftoff of the PMMA resist layer. The pattern has been written in the electronbeam writer, developed, and the metal layers deposited; however, the unpatterned glass surface is still covered with the PMMA bilayer, with metal on top. The large 1 mm square registers are visible, with sub-50-nm holes in a square lattice with a 2 mm unit cell spacing in between.

Palma et al.

3.1.6. Liftoff

Liftoff is the process to remove the remaining, unexposed resist that is now covered with metal. Recall that a slight overhang was created due to the use of a bilayer. Therefore, metals deposited inside the features on the glass substrate are not connected to the bulk of the metal sitting on top of the resist. Solvent is used to dissolve the resist, removing the metal while leaving behind only the metal deposited in the holes (directly on the glass) as defined by beam writing. The metal will appear to float off of the surface, hence the term liftoff. 1. To remove the resist, place the metal-coated sample in spectroscopic grade acetone (see Note 15). 2. Seal the container with parafilm tightly to avoid evaporation, and let sit overnight to remove the resist and metal layer on top (see Note 16). 3. When the sample is visibly clean of the metal layer, transfer it to a fresh beaker of acetone. 4. Remove the sample from the acetone, and rinse first with acetone from a squirt bottle, and then with ethanol from a squirt bottle (see Note 17). 5. Finally, blow dry with a stream of inert gas (Ar or N2). The Au/Pd nanopatterns on glass are now ready for surface functionalization. Figure  2 displays an atomic force microscopy (AFM) image and profile of the sub-50-nm dots fabricated on a glass coverslip.

500 nm Line profile 2.5 2 1.5 1 0.5 0

0

47.548

200

0.103

0.764 nm

176

nm

400

600

Fig. 2. Atomic force microscopy image of the Au/Pd nanodots array on glass. Imaging was performed with a XE-100, advanced scanning probe microscope (PSIA). The image has been analyzed with the XEI software, version 1.6.

Controlled Confinement of DNA at the Nanoscale

177

Fig. 3. Scheme of the main steps of the bio-functionalization procedure.

3.2. Functionalization of Nanopatterned Surfaces

Figure  3 schematically displays the main steps of the bio-­ functionalization procedure. Starting with a nanopatterned surface, i.e., Au/Pd nanodots on a glass slide, carry out the following steps: 1. Prepare fresh piranha solution (see above). 2. Clean the nanopatterned surface slide by immersion for 3 min in 1 h 30 min-aged piranha solution (if piranha is used as freshly prepared, it will eat away the metal). The immersion can be done using a Teflon rack to hold the sample/s in the container. 3. Take the sample out of the piranha solution with clean tweezers and rinse in DI water for at least 5 min. 4. Rinse the sample with ethanol and blow dry with a gentle stream of inert gas (Ar or N2). 5. Place the dry sample in a plasma cleaner for 5 min at 18 W. 6. Prepare a 1 mM anhydrous ethanol solution of HS–(CH2)11– (C2H6O2)3–OH and HS–(CH2)11–(C2H6O2)3–biotin: mix at 3:1 ratio (see Note 18). 7. Pull the sample out of the plasma cleaner and immediately incubate (see Note 19) it in the freshly prepared EG/biotin– thiol solution (1.5 mL per 6-well plate is sufficient to incubate the glass slide of the dimension used here). Seal the 6-well plate container with parafilm, cover with Al foil, and incubate on a shaker overnight (12–18 h). In this way, a SAM exhibiting functional biotin end groups will be formed on the metal nanodots; such biotin groups can then be used to immobilize streptavidin on the dots.

178

Palma et al.

8. Prepare a fresh solution of 2  mg of mPEG 5000-silane in 25 mL of anhydrous toluene and add 30 mL of acetic acid (as a catalyst) (see Notes 20 and 21). 9. Rinse the sample in ethanol and blow dry with a gentle stream of inert gas (Ar or N2). 10. Place the dry sample in the PEG solution. Incubate in a glass container, sealed with parafilm and covered with Al foil for at least 24 h (up to 48 h). This step is fundamental to assure proper passivation of the glass surface surrounding the dots: the formation of a PEG layer prevents (or at least minimizes) any nonspecific protein and DNA adsorption from taking place. 11. When PEGylation is done, rinse the glass slide with acetone and then ethanol, and blow dry with Ar or N2. 12. Prepare a solution of PBS with 10 mg/mL of streptavidin and 1  mg/mL of albumin in PBS (see Note 22). Place the dry slide in 2 mL of such solution, seal the container with Parafilm, cover with Al foil, and incubate on the shaker at RT for 2 h (see Note 23). 13. Rinse the sample thoroughly with PBS (do not let the sample dry). Then, move the sample to a fresh well filled with PBS and incubate it for at least 30 min, while storing it foiled, on a shaker (see Note 24). 14. Meanwhile, prepare a solution of biotinylated DNA (see Note 25). For biotinylated dsDNA, prepare a solution of two complementary strands of 2 mM in TPBS at RT, one biotinylated at the 5¢ end and its complement labeled with a fluorophore (Cy3 in our case) at the 5¢ end. Heat the solution gradually (steps of approximately 2°C per min) to a temperature of 65°C; leave the solution at this temperature for 15 min. Then ramp the temperature up to 75°C and leave for 1 h 30 min. Then proceed in the reverse order, ramping down the temperature, let cool at 65°C for 15 min and then take gradually to RT. When the solution has reached RT, it can be employed as described in the next step. 15. Incubate the (wet) coverslip from step 13, in at least 1.5 mL of the 2  mM biotinylated dsDNA solution prepared as described above (see Note 25); cover the container with Al foil during the incubation and seal with Parafilm. The minimum incubation time recommended is 3 h (see Note 26). 16. Take the sample out of the previous solution and rinse it with PBS. Then move the sample to a fresh well and incubate it for 30 min in PBS, while storing it foiled, on a shaker. The sample is now ready: i.e., Au/Pd nanodot is properly and specifically functionalized with the oligonucleotide of interest.

Controlled Confinement of DNA at the Nanoscale

179

In the next section, we will describe how we characterized the functionalized surface, verifying by fluorescence microscopy the presence of Cy3-labeled dsDNA immobilized on sub-50-nm Au/Pd nanodots on glass. 3.3. Fluorescence Microscopy 3.3.1. Sample Preparation

Inverted microscopes are designed to accept coverslips mounted on standard glass slides. We have devised a simple mounting scheme with our coverslips for fluorescence microscopy (see Fig. 4). It is an open design, which allows for additional processes to be carried out in situ on the microscope. Microfluidics can and have been used: here, we present a setup offering most of the functionality without the complexity of microfluidics. 1. Remove the bio-functionalized sample from PBS rinse solution and place face up on a clean surface. 2. Apply vacuum grease to the edge of a cloning ring and place it on the coverslip, with the pattern centered as well as can be done by hand. The grease creates a watertight seal. 3. Fill the ring with buffer solution (PBS). At this point, check for leaks around the ring. If any are detected, a slight twist of the ring is often adequate to seal the leak; if not, additional grease applied to the region of the leak will solve the problem. 4. Suspend the slide across the cutout section of the aluminum holder of the microscope. We use an aluminum plate cut to fit

Fig. 4. Photograph (Nikon D80) of the fully functionalized nanopatterned slide prepared for fluorescence microscopy. The coverslip sits atop an aluminum carrier, machined to fit the microscope mounting hardware, and is suspended above a cutout section allowing clearance for the optics. It is held in place by a thin bead of grease along each edge. A cloning ring adhered with a thin bead of grease encircles the patterned region and contains the necessary buffer solution.

180

Palma et al.

the microscope mount, with a gap cut into it to allow for the objective. 5. Use vacuum grease to seal the coverslip to the aluminum holder along the edges. Proceed to the microscope for imaging. 3.3.2. Microscopy

An inverted fluorescence microscope capable of epifluorescence microscopy is used to image the samples. Oil immersion lenses with 60× and 100× magnification are best used for imaging of the nanopatterns. The camera we use is a photometrics Cascade II; it is cooled to −70°C and has on-chip amplification for low noise, high-sensitivity imaging. 1. Make sure the optics are aligned. 2. Find the pattern in differential image contrast (DIC) mode. 3. Switch from DIC to the Cy3 (excitation 550 nm/emission 568 nm) fluorescence channel. Use live imaging with