M e t h o d s i n M o l e c u l a r B i o l o g y ™

Series EditorJohn M. Walker

School of Life SciencesUniversity 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

EditorsGiampaolo Zuccheri, Ph.D.Department of Biochemistry University of Bologna Bologna, Italygiampaolo.zuccheri@unibo.it

Bruno SamorìDepartment of Biochemistry University of Bologna Bologna, Italybruno.samori@unibo.it

ISSN 1064-3745 e-ISSN 1940-6029ISBN 978-1-61779-141-3 e-ISBN 978-1-61779-142-0DOI 10.1007/978-1-61779-142-0Springer New York Heidelberg London Dordrecht

Library of Congress Control Number: 2011929163

© Springer Science+Business Media, LLC 2011All 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.

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v

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”) refer-ring 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 opportuni-ties 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 proper-ties 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 proto-cols. Second, the DNA molecule can be decontextualized and “simply” used as a copoly-mer 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: every-thing 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 nano-object, such as a DNA origami, you marvel at how smart DNA is as a molecule and won-der 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

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 nanostruc-tures, 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 nanostruc-tures 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 pro-tocols 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 automati-cally, possibly with a precision and output rate similar to that of the assembly of microelec-tronics circuitry nowadays.

Our personal wish is that the next chapters will be written by some of our readers.

Bologna, Italy Giampaolo ZuccheriBologna, Italy Bruno Samorì

vii

Contents

Preface. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . vContributors . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . ix

1 Synthesis and Characterization of Self-Assembled DNA Nanostructures . . . . . . . . 1Chenxiang Lin, Yonggang Ke, Rahul Chhabra, Jaswinder Sharma, Yan Liu, and Hao Yan

2 Protocols for Self-Assembly and Imaging of DNA Nanostructures . . . . . . . . . . . . 13Thomas L. Sobey and Friedrich C. Simmel

3 Self-Assembly of Metal-DNA Triangles and DNA Nanotubes with Synthetic Junctions . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 33Hua Yang, Pik Kwan Lo, Christopher K. McLaughlin, Graham D. Hamblin, Faisal A. Aldaye, and Hanadi F. Sleiman

4 DNA-Templated Pd Conductive Metallic Nanowires . . . . . . . . . . . . . . . . . . . . . . 49Khoa Nguyen, Stephane Campidelli, and Arianna Filoramo

5 A Method to Map Spatiotemporal pH Changes Inside Living Cells Using a pH-Triggered DNA Nanoswitch. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 61Souvik Modi and Yamuna Krishnan

6 Control of Helical Handedness in DNA and PNA Nanostructures . . . . . . . . . . . . 79Roberto Corradini, Tullia Tedeschi, Stefano Sforza, Mark M. Green, and Rosangela Marchelli

7 G-Quartet, G-Quadruplex, and G-Wire Regulated by Chemical Stimuli . . . . . . . . 93Daisuke Miyoshi and Naoki Sugimoto

8 Preparation and Atomic Force Microscopy of Quadruplex DNA . . . . . . . . . . . . . . 105James Vesenka

9 Synthesis of Long DNA-Based Nanowires. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 115Alexander Kotlyar

10 G-Wire Synthesis and Modification with Gold Nanoparticle . . . . . . . . . . . . . . . . . 141Christian Leiterer, Andrea Csaki, and Wolfgang Fritzsche

11 Preparation of DNA Nanostructures with Repetitive Binding Motifs by Rolling Circle Amplification . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 151Edda Reiß, Ralph Hölzel, and Frank F. Bier

12 Controlled Confinement of DNA at the Nanoscale: Nanofabrication and Surface Bio-Functionalization . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 169Matteo 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 . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 187Albert M. Hung and Jennifer N. Cha

14 DNA-Modified Single Crystal and Nanoporous Silicon. . . . . . . . . . . . . . . . . . . . . 199Andrew Houlton, Bernard A. Connolly, Andrew R. Pike, and Benjamin R. Horrocks

viii Contents

15 The Atomic Force Microscopy as a Lithographic Tool: Nanografting of DNA Nanostructures for Biosensing Applications . . . . . . . . . . . . . . . . . . . . . . . 209Matteo Castronovo and Denis Scaini

16 Trapping and Immobilization of DNA Molecules Between Nanoelectrodes. . . . . . 223Anton Kuzyk, J. Jussi Toppari, and Päivi Törmä

17 DNA Contour Length Measurements as a Tool for the Structural Analysis of DNA and Nucleoprotein Complexes. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 235Claudio Rivetti

18 DNA Molecular Handles for Single-Molecule Protein-Folding Studies by Optical Tweezers . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 255Ciro 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. . . . . . . . . . . . . . . . . 273Matt V. Fagerburg and Sanford H. Leuba

20 Engineering Mononucleosomes for Single-Pair FRET Experiments. . . . . . . . . . . . 291Wiepke J.A. Koopmans, Ruth Buning, and John van Noort

21 Measuring DNA–Protein Binding Affinity on a Single Molecule Using Optical Tweezers . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 305Micah J. McCauley and Mark C. Williams

22 Modeling Nanopores for Sequencing DNA. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 317Jeffrey R. Comer, David B. Wells, and Aleksei Aksimentiev

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

ix

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, GermanymAtt 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, GermanyAlon A. gorodetsky • Department of Chemistry, Columbia University,

New York, NY, USA

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, GermanysAnFord 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, CanadarosAngelA mArChelli • Dipartimento di Chimica Organica e Industriale,

Univeristà di Parma, Parma, ItalysusAn mArqusee • Department of Molecular & Cell Biology, University of

California, Berkeley, CA, USAmiCAh J. mCCAuley • Department of Physics, Northeastern University,

Boston, MA, USAChristopher k. mClAughlin • Department of Chemistry, McGill University,

Montreal, CanadadAisuke miyoshi • Faculty of Frontiers of Innovative Research in Science

and Technology (FIRST), and Frontier Institute for Biomolecular Engineering Research (FIBER), Konan University, Kobe, Japan

xiContributors

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, ItalysteFAno sForzA • Dipartimento di Chimica Organica e Industriale,

Univeristà di Parma, Parma, ItalyelizAbeth A. shAnk • Harvard Medical School, Boston, MA, USAJAsWinder shArmA • Center for Integrated Nanotechnologies, Los Alamos

National Laboratory, Los Alamos, NM, USAmiChAel p. sheetz • Department of Biological Sciences, Columbia University,

New York, NY, USAFriedriCh C. simmel • Physik Department, Technische Universität München,

Munich, GermanyhAnAdi F. sleimAn • Department of Chemistry, McGill University, Montreal,

CanadathomAs l. sobey • Physik Department, Technische Universität München,

Munich, GermanynAoki 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

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

105

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

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 self-assembly 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 atmo-sphere imaging conditions. Detailed step-wise instructions are provided.

Key words: G-wire, Quadruplex DNA, Guanine-rich oligonucleotide, Self-assembly, Atomic force microscopy

G-DNA is a polymorphic family of four-stranded structures con-taining 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 resi-dues 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

1. Introduction

106 Vesenka

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.

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 ).

107Preparation and Atomic Force Microscopy of Quadruplex DNA

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.

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 neighbor-hood of 100 mM, but must be diluted for AFM imaging of individual G-wires.

2. Materials

G GK+

Mg2+ M+

b

N NcG G- -

--

--

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 unsup-ported phosphate backbone of duplex DNA, shown here as N–N

c (complementary) base pair, electrostatically pins the duplex DNA flat to the mica substrate.

108 Vesenka

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 syntheti-cally 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).

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 substitut-ing 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].

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

2.1. GRO Self-assembly

2.2. Imaging Buffers

3. Methods

3.1. GRO Self-assembly

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.

109Preparation and Atomic Force Microscopy of Quadruplex DNA

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 dis-assembled 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, evapora-tion of buffer can present a problem. Evaporation can be reme-died 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 tempera-ture, 4°C, or freezing appears to have no impact on G-wire integrity.

1. Attach a mica disk (high-quality, 10 mm diameter disks were used in this work) to a ferrous disk (steel “puck”) using adhe-sive 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 accom-plished 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 consid-ered: 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

3.2. AFM Sample Preparation and Imaging

111Preparation and Atomic Force Microscopy of Quadruplex DNA

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 con-trolled (e.g., NDT), and other systems require “false engage-ment” (e.g., Digital Instruments/Veeco) to avoid tip contact (see Note 6).

10. Humidity and thermal equilibrium can be achieved by install-ing 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 micro-scope to equilibrate for several hours while electronic stabili-zation 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 pro-portionally increased with decrease in scan size. The impor-tance 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 auto-matically 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 engage-ment value and capture the image.

1. The monovalent cation stabilizes the G-quartet. The magne-sium ions and spermidine stabilize the G-wire phosphate back-bone. Prepare buffer from fresh deionized water at 10× the

4. Notes

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):

= » m1 A unit ( 1OD) of single-stranded DNA 33 g/mL260

#( g) 3.0 (nmole/( g)/N #(nmole),m ´ m =

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 evapo-rate 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 deion-ized 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 prepara-tion 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 scan-ner to engage falsely and scan at the speed and size deter-mined 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.

113Preparation and Atomic Force Microscopy of Quadruplex DNA

References

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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 oli-gomers. 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 micros-copy, 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 biomole-cules 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 gold-labeled 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 nano-tube 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.