Single-Molecule Enzymology: Nanomechanical Manipulation and Hybrid Methods [1st Edition] 9780128095034, 9780128093108

Table of contents :
Content:
Series PagePage ii
CopyrightPage iv
ContributorsPages xi-xiv
PrefacePages xv-xviMaria Spies, Yann R. Chemla
Chapter One - How to Measure Load-Dependent Kinetics of Individual Motor Molecules Without a Force-ClampPages 1-29J. Sung, K.I. Mortensen, J.A. Spudich, H. Flyvbjerg
Chapter Two - Studying the Mechanochemistry of Processive Cytoskeletal Motors With an Optical TrapPages 31-54V. Belyy, A. Yildiz
Chapter Three - Single-Molecule Optical-Trapping Techniques to Study Molecular Mechanisms of a ReplisomePages 55-84B. Sun, M.D. Wang
Chapter Four - Recent Advances in Biological Single-Molecule Applications of Optical Tweezers and Fluorescence MicroscopyPages 85-119M. Hashemi Shabestari, A.E.C. Meijering, W.H. Roos, G.J.L. Wuite, E.J.G. Peterman
Chapter Five - Direct Visualization of Helicase Dynamics Using Fluorescence Localization and Optical TrappingPages 121-136C.-T. Lin, T. Ha
Chapter Six - High-Resolution Optical Tweezers Combined With Single-Molecule Confocal MicroscopyPages 137-169K.D. Whitley, M.J. Comstock, Y.R. Chemla
Chapter Seven - Integrating Optical Tweezers, DNA Tightropes, and Single-Molecule Fluorescence Imaging: Pitfalls and TrapsPages 171-192J. Wang, J.T. Barnett, M.R. Pollard, N.M. Kad
Chapter Eight - Single-Stranded DNA Curtains for Studying Homologous RecombinationPages 193-219C.J. Ma, J.B. Steinfeld, E.C. Greene
Chapter Nine - Inserting Extrahelical Structures into Long DNA Substrates for Single-Molecule Studies of DNA Mismatch RepairPages 221-238M.W. Brown, A. de la Torre, I.J. Finkelstein
Chapter Ten - Single-Molecule Insight Into Target Recognition by CRISPR–Cas ComplexesPages 239-273M. Rutkauskas, A. Krivoy, M.D. Szczelkun, C. Rouillon, R. Seidel
Chapter Eleven - Preparation of DNA Substrates and Functionalized Glass Surfaces for Correlative Nanomanipulation and Colocalization (NanoCOSM) of Single MoleculesPages 275-296C. Duboc, J. Fan, E.T. Graves, T.R. Strick
Chapter Twelve - Measuring Force-Induced Dissociation Kinetics of Protein Complexes Using Single-Molecule Atomic Force MicroscopyPages 297-320K. Manibog, C.F. Yen, S. Sivasankar
Chapter Thirteen - Improved Force Spectroscopy Using Focused-Ion-Beam-Modified CantileversPages 321-351J.K. Faulk, D.T. Edwards, M.S. Bull, T.T. Perkins
Chapter Fourteen - Single-Molecule Characterization of DNA–Protein Interactions Using Nanopore BiosensorsPages 353-385A.H. Squires, T. Gilboa, C. Torfstein, N. Varongchayakul, A. Meller
Chapter Fifteen - Subangstrom Measurements of Enzyme Function Using a Biological Nanopore, SPRNTPages 387-414A.H. Laszlo, I.M. Derrrington, J.H. Gundlach
Chapter Sixteen - Multiplexed, Tethered Particle Microscopy for Studies of DNA-Enzyme DynamicsPages 415-435S. Ucuncuoglu, D.A. Schneider, E.R. Weeks, D. Dunlap, L. Finzi
Author IndexPages 437-458
Subject IndexPages 459-467

Citation preview

METHODS IN ENZYMOLOGY Editors-in-Chief

ANNA MARIE PYLE Departments of Molecular, Cellular and Developmental Biology and Department of Chemistry Investigator, Howard Hughes Medical Institute Yale University

DAVID W. CHRISTIANSON Roy and Diana Vagelos Laboratories Department of Chemistry University of Pennsylvania Philadelphia, PA

Founding Editors

SIDNEY P. COLOWICK and NATHAN O. KAPLAN

Academic Press is an imprint of Elsevier 50 Hampshire Street, 5th Floor, Cambridge, MA 02139, United States 525 B Street, Suite 1800, San Diego, CA 92101–4495, United States The Boulevard, Langford Lane, Kidlington, Oxford OX5 1GB, United Kingdom 125 London Wall, London, EC2Y 5AS, United Kingdom First edition 2017 Copyright © 2017 Elsevier Inc. All rights reserved. No part of this publication may be reproduced or transmitted in any form or by any means, electronic or mechanical, including photocopying, recording, or any information storage and retrieval system, without permission in writing from the publisher. Details on how to seek permission, further information about the Publisher’s permissions policies and our arrangements with organizations such as the Copyright Clearance Center and the Copyright Licensing Agency, can be found at our website: www.elsevier.com/permissions. This book and the individual contributions contained in it are protected under copyright by the Publisher (other than as may be noted herein). Notices Knowledge and best practice in this field are constantly changing. As new research and experience broaden our understanding, changes in research methods, professional practices, or medical treatment may become necessary. Practitioners and researchers must always rely on their own experience and knowledge in evaluating and using any information, methods, compounds, or experiments described herein. In using such information or methods they should be mindful of their own safety and the safety of others, including parties for whom they have a professional responsibility. To the fullest extent of the law, neither the Publisher nor the authors, contributors, or editors, assume any liability for any injury and/or damage to persons or property as a matter of products liability, negligence or otherwise, or from any use or operation of any methods, products, instructions, or ideas contained in the material herein. ISBN: 978-0-12-809310-8 ISSN: 0076-6879 For information on all Academic Press publications visit our website at https://www.elsevier.com/

Publisher: Zoe Kruze Acquisition Editor: Zoe Kruze Editorial Project Manager: Helene Kabes Production Project Manager: Magesh Kumar Mahalingam Cover Designer: Greg Harris Typeset by SPi Global, India

CONTRIBUTORS J.T. Barnett School of Biosciences, University of Kent, Canterbury, Kent, United Kingdom V. Belyy Biophysics Graduate Group, University of California at Berkeley, Berkeley, CA, United States M.W. Brown Institute for Cellular and Molecular Biology, The University of Texas at Austin, Austin, TX, United States M.S. Bull JILA, National Institute of Standards and Technology and University of Colorado, Boulder, CO, United States Y.R. Chemla Center for the Physics of Living Cells; University of Illinois at Urbana–Champaign, Urbana, IL, United States M.J. Comstock Michigan State University, East Lansing, MI, United States A. de la Torre Institute for Cellular and Molecular Biology, The University of Texas at Austin, Austin, TX, United States I.M. Derrrington University of Washington, Seattle, WA, United States C. Duboc Institut Jacques Monod, Centre National de la Recherche Scientifique, University of Paris Diderot and Sorbonne Paris Cit
e, Paris, France D. Dunlap Emory University, Atlanta, GA, United States D.T. Edwards JILA, National Institute of Standards and Technology and University of Colorado, Boulder, CO, United States J. Fan Institut Jacques Monod, Centre National de la Recherche Scientifique, University of Paris Diderot and Sorbonne Paris Cit
e, Paris, France J.K. Faulk JILA, National Institute of Standards and Technology and University of Colorado, Boulder, CO, United States

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Contributors

I.J. Finkelstein Institute for Cellular and Molecular Biology; Center for Systems and Synthetic Biology, The University of Texas at Austin, Austin, TX, United States L. Finzi Emory University, Atlanta, GA, United States H. Flyvbjerg Department of Micro- and Nanotechnology, Technical University of Denmark, Kongens Lyngby, Denmark T. Gilboa The Technion, Haifa, Israel E.T. Graves Institut Jacques Monod, Centre National de la Recherche Scientifique, University of Paris Diderot and Sorbonne Paris Cit
e, Paris, France E.C. Greene Columbia University, New York, NY, United States J.H. Gundlach University of Washington, Seattle, WA, United States M. Hashemi Shabestari Vrije Universiteit, Amsterdam, The Netherlands T. Ha Johns Hopkins University; Howard Hughes Medical Institute, Baltimore, MD, United States N.M. Kad School of Biosciences, University of Kent, Canterbury, Kent, United Kingdom A. Krivoy Molecular Biophysics Group, Institute for Experimental Physics I, Universit€at Leipzig, Leipzig, Germany; Skolkovo Institute of Science and Technology, Skolkovo, Russia A.H. Laszlo University of Washington, Seattle, WA, United States C.-T. Lin Johns Hopkins University, Baltimore, MD, United States C.J. Ma Columbia University, New York, NY, United States K. Manibog Iowa State University; Ames Laboratory, U.S. Department of Energy, Ames, IA, United States A.E.C. Meijering Vrije Universiteit, Amsterdam, The Netherlands A. Meller The Technion, Haifa, Israel; Boston University, Boston, MA, United States

Contributors

xiii

K.I. Mortensen Department of Micro- and Nanotechnology, Technical University of Denmark, Kongens Lyngby, Denmark T.T. Perkins JILA, National Institute of Standards and Technology; University of Colorado, Boulder, CO, United States E.J.G. Peterman Vrije Universiteit, Amsterdam, The Netherlands M.R. Pollard DFM A/S, Kongens Lyngby, Denmark W.H. Roos Moleculaire Biofysica, Zernike Institute, Rijksuniversiteit Groningen, Groningen, The Netherlands C. Rouillon Molecular Biophysics Group, Institute for Experimental Physics I, Universit€at Leipzig, Leipzig, Germany M. Rutkauskas Molecular Biophysics Group, Institute for Experimental Physics I, Universit€at Leipzig, Leipzig, Germany D.A. Schneider University of Alabama at Birmingham, Birmingham, AL, United States R. Seidel Molecular Biophysics Group, Institute for Experimental Physics I, Universit€at Leipzig, Leipzig, Germany S. Sivasankar Iowa State University; Ames Laboratory, U.S. Department of Energy, Ames, IA, United States J.A. Spudich Department of Biochemistry, Stanford University School of Medicine, Stanford, CA, United States A.H. Squires Stanford University, Stanford, CA, United States J.B. Steinfeld Columbia University, New York, NY, United States T.R. Strick Institut Jacques Monod, Centre National de la Recherche Scientifique, University of Paris Diderot and Sorbonne Paris Cit
e; Ecole Normale Sup
erieure, Institut de Biologie de l’ENS (iBENS), INSERM, CNRS, PSL Research University, Paris, France B. Sun School of Life Science and Technology, ShanghaiTech University, Shanghai, PR China

xiv

Contributors

J. Sung Department of Cellular and Molecular Pharmacology, The Howard Hughes Medical Institute, University of California, San Francisco, CA, United States M. Szczelkun DNA–Protein Interactions Unit, School of Biochemistry, University of Bristol, Bristol, United Kingdom C. Torfstein The Technion, Haifa, Israel S. Ucuncuoglu Emory University, Atlanta, GA, United States N. Varongchayakul Boston University, Boston, MA, United States J. Wang School of Biosciences, University of Kent, Canterbury, Kent, United Kingdom M.D. Wang Laboratory of Atomic and Solid State Physics; Howard Hughes Medical Institute, Cornell University, Ithaca, NY, United States E.R. Weeks Emory University, Atlanta, GA, United States K.D. Whitley University of Illinois at Urbana–Champaign, Urbana, IL, United States G.J.L. Wuite Vrije Universiteit, Amsterdam, The Netherlands C.F. Yen Iowa State University; Ames Laboratory, U.S. Department of Energy, Ames, IA, United States A. Yildiz University of California at Berkeley, Berkeley, CA, United States

PREFACE Single-molecule biophysics is a burgeoning field that encompasses studies of individual molecules and macromolecular machines in isolation, within the cells, and even inside living organisms. The last two decades witnessed an explosion of single-molecule research. The technological advances made to visualize and investigate biologically important molecules individually, in real time, and under physiological conditions have fundamentally changed our understanding of how cells work and communicate, how the molecular machines of the cell assemble and function, and, in general, how physiological functions emerge from the chaos of stochastic molecular events and interactions. New insights have been gained into the mechanisms of molecular motors, cell division, DNA repair and replication, RNA transcription, translation, protein folding, enzymatic catalysis, and assembly and function of membrane proteins. The time when the novelty of observing a single molecule was a cause for celebration has passed. Single-molecule techniques have grown more sensitive and sophisticated, capable of multidimensional or high-throughput measurements of biomolecular dynamics. Once arcane, single-molecule methodologies are also becoming more mainstream. With the technologies becoming increasingly available to a broader community of biophysicists, biochemists, and molecular biologists, what lags behind is a comprehensive understanding of the power of single-molecule techniques when combined with rigorous data analysis. Thus, we believe the time is right for a comprehensive text describing the most cutting-edge single-molecule methods. The two volumes in this “Single-Molecule Enzymology” set are not intended to replace the previous set (volumes 472 and 475 on “SingleMolecule Tools”)—all the methods and analyses described in it are still highly relevant. Instead, our aim is to expand the previous volumes by providing a comprehensive toolkit of up-to-date methods and expert advice on their utilization and data analysis. The first volume in the set (volume 581, “Single-Molecule Enzymology, Part A”) focuses on advances in the fluorescence-based techniques including single-molecule fluorescence in single or multiple colors, FRET, multidimensional FRET, interferometry, and massively parallel single-molecule enzymatic analyses in nanoliter volumes. The second volume in the series (volume 582, “Single-Molecule Enzymology, Part B”) focuses on the xv

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force-based single-molecule techniques and hybrid approaches that combine manipulation of single molecules by force with fluorescence-based observation. Our philosophy when we selected the topics and contributors was to offer an authoritative practical guide on the most popular and newly emerging aspects of single-molecule enzymology. Our hope is that the two volumes in this series will provide valuable information on everything that goes into a successful single-molecule experiment from sample preparation, to measurements, data acquisition, and rigorous data analysis routines, and will be well used by everyone in the lab, from undergraduates and rotation students to postdoctoral fellows and faculty who wish to expand their research programs. MARIA SPIES University of Iowa YANN R. CHEMLA University of Illinois, Urbana-Champaign

CHAPTER ONE

How to Measure Load-Dependent Kinetics of Individual Motor Molecules Without a Force-Clamp J. Sung*, K.I. Mortensen†, J.A. Spudich{, H. Flyvbjerg†,1 *Department of Cellular and Molecular Pharmacology, The Howard Hughes Medical Institute, University of California, San Francisco, CA, United States † Department of Micro- and Nanotechnology, Technical University of Denmark, Kongens Lyngby, Denmark { Department of Biochemistry, Stanford University School of Medicine, Stanford, CA, United States 1 Corresponding author: e-mail address: [email protected]

Contents 1. Introduction 1.1 Single-Molecule Force Spectroscopy Techniques 1.2 Force-Clamp Optical Trap 1.3 Harmonic Force Spectroscopy 2. HFS: Basic Concept 3. Experimental Setup 4. Sample Preparations: Proteins, Reagents, and Buffers 4.1 Human β-Cardiac Myosin S1 4.2 Actin Filaments 4.3 Anti-GFP Antibody 4.4 Trapping Beads 4.5 Nitrocellulose-Coated Coverslip 4.6 Other Reagents 4.7 Assay Buffer (AB) 4.8 AB with BSA (ABBSA) 4.9 GO buffer 5. Experimental Protocols 5.1 Preparation of Flow-Cell Sample Chamber 5.2 Formation of an Actin Dumbbell 5.3 HFS: Experiment 6. Trap Calibration 7. HFS: Theory and Data Analysis 7.1 Automatic Binding Detection 7.2 Force-Dependent Kinetics Under Harmonic Force 7.3 Mathematical Model and Theory 8. Results and Discussion 9. Conclusion and Outlook Acknowledgments References Methods in Enzymology, Volume 582 ISSN 0076-6879 http://dx.doi.org/10.1016/bs.mie.2016.08.002

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2017 Elsevier Inc. All rights reserved.

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Abstract Single-molecule force spectroscopy techniques, including optical trapping, magnetic trapping, and atomic force microscopy, have provided unprecedented opportunities to understand biological processes at the smallest biological length scales. For example, they have been used to elucidate the molecular basis of muscle contraction and intracellular cargo transport along cytoskeletal filamentous proteins. Optical trapping is among the most sophisticated single-molecule techniques. With exceptionally high spatial and temporal resolutions, it has been extensively utilized to understand biological functions at the single molecule level, such as conformational changes and forcegeneration of individual motor proteins or force-dependent kinetics in molecular interactions. Here, we describe a new method, “Harmonic Force Spectroscopy (HFS).” With a conventional dual-beam optical trap and a simple harmonic oscillation of the sample stage, HFS can measure the load-dependent kinetics of transient molecular interactions, such as a human β-cardiac myosin II interacting with an actin filament. We demonstrate that the ADP release rate of an individual human β-cardiac myosin II molecule depends exponentially on the applied load, which provides a clue to understanding the molecular mechanism behind the force–velocity curve of a contracting cardiac muscle. The experimental protocol and the data analysis are simple, fast, and efficient. This chapter provides a practical guide to the method: basic concepts, experimental setup, step-bystep experimental protocol, theory, data analysis, and results.

1. INTRODUCTION 1.1 Single-Molecule Force Spectroscopy Techniques During the past two decades, significant progress has been made in understanding the molecular basis of load-dependence, primarily due to developments of new methodologies that are capable of directly manipulating forces and characterizing force-dependent functional changes at the molecular level. In particular, various single-molecule force spectroscopy techniques, including optical trapping, magnetic trapping, and atomic force microscopy, have been developed and utilized in characterizing detailed mechanisms of load-dependence at the single-molecule level (Capitanio & Pavone, 2013; Greenleaf, Woodside, & Block, 2007; Neuman & Nagy, 2008). Optical trapping, in particular, is among the most sophisticated single-molecule force spectroscopy techniques with exceptionally high spatial (nanometers) and temporal (milliseconds) resolution, which can directly characterize conformational changes, binding/unbinding kinetics, and force-dependent enzyme kinetics (Capitanio & Pavone, 2013; Neuman & Block, 2004).

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1.2 Force-Clamp Optical Trap Force-clamp is one configuration of an optical trap that can directly measure force-dependent kinetics or conformational changes of proteins or nucleic acids via sophisticated active feedback control of the trap. For example, Veigel et al. showed that the lifetime of the bound crossbridge between a smooth-muscle myosin and an actin filament is modulated by an applied load, which provides a clue in explaining the “Fenn effect” in muscle contraction (Veigel, Molloy, Schmitz, & Kendrick-Jones, 2003). A conventional force-clamp is usually operated in the following steps: (1) detect the event, such as binding or conformational changes; (2) apply a constant force by an active feedback control of the trap setup; (3) measure the lifetime of the state at the applied constant force; and (4) repeat the previous steps to obtain enough data at the applied force. The measurement is performed multiple times at different forces to obtain the force-dependent kinetics of the molecule. Success of a force-clamp experiment is mainly determined by (i) the speed and accuracy with which the true signal is detected in the presence of noise and (ii) the speed of the ensuing feedback response of the trap, working in a highly reliable manner. The noise consists of intrinsic Brownian noise and instrument noise. Speed and accuracy, however, compete with each other. For example, in order to improve accuracy, one needs more data, longer time series, before deciding about detection, which in turn delays the feedback response time. This can be especially problematic if the signal of interest, such as a binding lifetime or a conformational state, is transient and similar to or shorter than the time needed for reliable detection and feedback response. Instead, one can use shorter time series, less data in order to speed up the detection, but that inevitably increases the chance of errors, such as false-positive or false-negative events. Therefore, it has been the main focus in the field to develop improved feedback algorithms that work both quickly and accurately. One such example is the ultrafast forceclamp technique recently developed by Capitanio et al. (2012). Oscillating the trap in a triangular waveform and fast feedback control of the trap increased the detection sensitivity and the speed, achieving sub-ms temporal resolution. A drawback of the active feedback methods is that they all require a complicated apparatus for fast and accurate on-line detection of the events, followed by careful control of the system for robust operation of the feedback setup, which is technically challenging. In addition, this complication

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for the operation makes data sampling less efficient and hence lowers throughput, making it difficult to collect enough data to assess the loaddependence of individual molecules. Thus, earlier methods (Greenberg, Shuman, & Ostap, 2014; Veigel et al., 2003) average over results from multiple molecules, though obtained individually, to characterize the force dependence, based on the assumption that the molecules all are identical, which may or may not be true.

1.3 Harmonic Force Spectroscopy We recently developed an approach orthogonal to current force-clamp methods, a new method that we refer to as harmonic force spectroscopy (HFS) for reasons given later (Sung et al., 2015). HFS can detect and apply forces to weak and transient molecular interactions. Its experimental protocol and data analysis are simple, fast, and efficient. HFS runs automatically, without any feedback control, and applies a randomly chosen force to a single motor protein, practically without delay, after the motor protein has bound to its track. Moreover, the sampling efficiency is improved with HFS, so we could collect enough data from an individual molecule for a range of forces to get full load-dependent kinetics curves for individual molecules. In Sung et al. (2015), we demonstrated the power of HFS by directly measuring the force-dependent ADP release kinetics of human β-cardiac myosin II (Fig. 1), as an example of load-dependent transient molecular interactions. The method, however, is not limited to actin–myosin interactions, but can be modified and applied to other systems where force matters in molecular interactions, such as microtubule-associated proteins (MAPs including kinesin and dynein motors) interacting with a microtubule track, or DNA/RNA-binding proteins interacting with their filamentous track. In this chapter, we provide a practical guide to an experiment using HFS. We describe the basic concept of HFS, the experimental setup, preparations of samples and reagents, the step-by-step experimental protocol to obtain data, the theory and data analysis, and conclude with conclusion and outlook.

2. HFS: BASIC CONCEPT Here, we briefly describe the basic concept of the method before we get into the details. The way it works is quite simple. We trap two beads using a dual-beam trap, and bridge them with an actin filament, called an actin dumbbell (details later). Motors are anchored to the surface, some of which are at the top of a platform-bead stuck to the surface. We move

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Fig. 1 Actin-activated ATPase cycle of human β-cardiac S1. The actin-activated ATPase cycle of myosin illustrates the conformational states and the nucleotide states of the motor in association with an actin filament. The actin filament is shown in gray, myosin is shown in red in the strongly bound states, and in yellow in the weakly bound states. Arrows indicate the transition between the states. At saturating ATP concentration, the rate limiting step of the overall strongly bound states is the ADP release step, which takes
10 ms with human β-cardiac S1. The time spent in the strongly bound state divided by the cycling time is the duty-ratio, which is
10% with human β-cardiac S1.

the dumbbell in proximity to the platform bead to let the surface-attached motor bind to the actin dumbbell (Fig. 2). The motor consequently undergoes a power-stroke immediately, which results in an abrupt displacement of the dumbbell. Binding of ATP to the motor terminates the myosin– actin bond, and the freed dumbbell goes back to its original position. This defines a single myosin–actin binding event, which repeats multiple times until the active motor becomes inactive. So far, this description is just that of the conventional dual-beam threebead assay for myosin, and we are now ready to start the HFS measurements. Now, we simply oscillate the sample sinusoidally, using a piezo-electric stage (PZT) (alternatively one may oscillate the dumbbell using an acousto-optic deflector (AOD) or acousto-optic modulator (AOM)), while the motor and the actin dumbbell undergo cycles of binding and unbinding (see Fig. 2 for a schematic illustration). When the motor binds and holds the dumbbell, the oscillatory translation of the motor with the stage is transferred to the position of the dumbbell, which results in harmonic oscillation of the dumbbell. This oscillation amplifies the binding signal and enables us to detect readily brief and weak binding events, which are difficult to detect without this or similar

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Fig. 2 Illustration of the harmonic force spectroscopy experiment. Two trapped beads (dark gray) are bridged with an actin filament (light gray) and tightly stretched by dualbeam traps. Myosin is anchored on a platform bead and harmonically oscillated by a piezo-electric stage. The platform bead is positioned in the middle of the dumbbell, slightly below the actin filament. Lowering the dumbbell close to the platform bead will allow the motor to bind to actin and oscillate the trapped beads. Binding can take place anywhere in the oscillation cycle. This figure is not to scale.

amplification. We continue the measurement to obtain multiple binding events from the same molecule until the motor becomes inactive. We then correlate the force applied to the motor and the time spent in the bound state. Since the binding occurs at different phases of the oscillation, a spectrum of different mean forces are applied, as the cycle-averaged force depends on where in the cycle binding took place. For example, the motor can bind when it is on the left side of the oscillation (Fig. 3A). In this case, the motor is under net negative (or backward) mean force. On the other hand, the binding site could be in the central region (Fig. 3B) or the right side of the oscillation (Fig. 3C), and different force will be applied accordingly. When we correlated the mean force and the dwell time, we observed that the human β-cardiac myosin S1 molecule displays highly asymmetric dwell time distribution, implying that the ADP release rate is force dependent. Note here that the experiment was done at saturating ATP concentration (2 mM). Unlike the case with a force-clamp, in HFS a motor experiences an oscillating force while bound. The quantitative description of this requires a little extra math, but it results in a force-dependent detachment rate that is identical to the one for constant force, except the mean force appears in the place of the constant force, and a “correction factor” appears, which describes the effect of force oscillations. Derived once and for all, we use this formula to obtain

t2

Δx (nm)

F0 ≈ 0

Distance (nm)

Zero load

Δx1

F

I

Distance (nm)

Δx (nm)

Time (ms)

C

t1

Forward load

F0 < 0

Δx1 Time (ms)

Fig. 3 See legend on next page.

Time (ms)

t1

K

Time (ms) ts t2

L

t2

x1(0) > 0

Time (ms) ts t2

Δx1

Distance (nm)

ts

Phase difference (rad)

t1

H

ts

Δx1

Time (ms)

Time (ms)

E

t1

Distance (nm)

Δx1

B

J

t2

ts

Phase difference (rad)

t1

x1(0) ≈ 0

t1

Time (ms) t2 ts

Distance (nm)

F0 > 0

G

Phase difference (rad)

Backward load

D

Δx (nm)

Center of the oscillation

Distance (nm)

A

x1(0) < 0 Δxx1

Time (ms)

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the force-dependent ADP release rate of individual human β-cardiac myosin S1 molecules under constant force. The rate we find fits well with the Bell– Evans model, describing bond rupture under force (Bell, 1978). HFS is so efficient that we could find the force-dependent ADP release rate of individual molecules. We did this for multiple molecules to test reproducibility and individuality of hypothetically identical molecules.

3. EXPERIMENTAL SETUP The experimental setup required for HFS in this study is a conventional dual-beam optical trap setup (Finer, Simmons, & Spudich, 1994; Sung, Sivaramakrishnan, Dunn, & Spudich, 2010) with capability of harmonic oscillation of the sample chamber via a PZT with a well-defined oscillation amplitude and frequency. Unlike other force-clamp methods, no feedback control of the trap is required to measure the load-dependent lifetime or kinetics. The detailed protocol on how to set up the dual-beam optical trap instrument used in this study is described in Sung et al. (2010). The instrumentation used in this study is described in detail elsewhere (Sung et al., 2015). Fig. 3 Myosin binding and unbinding under harmonic oscillation. Harmonic force spectroscopy with human β-cardiac S1. (A, B, C) An actin dumbbell held by two fixed optical traps interacts with a β-cardiac S1 molecule that is surface-attached to the top of a platform bead on a sinusoidally translating piezo-electric stage. Upon binding, the motor strokes toward the right (black arrows). Depending on whether the binding occurs left of the center (asterisk in (A)), at the center (asterisk on gray dashed line in (B)), or right of the center (asterisk in (C)), the motor experiences backward (A), near zero (B), or forward (C) mean load F0. (D, E, F) Three examples of time traces of the displacement x1 of a dumbbell bead in its trap. The other bead has coordinate x2 (not shown). Large sinusoidal oscillations caused by being attached to the stage by a motor are observed in the middle portion of the traces. Small oscillations before and after attachments are caused by drag from buffer moving past the unattached dumbbell. (G, H, I) Change in phase shift φ1(t) relative to the unbound state (black lines, dashed black lines showing its time average) and amplitude △x1 ðtÞ of oscillations (alternatively colored lines, dashed alternatively colored lines showing its time average obtained from longer stretches of unbound state times than shown) of bead positions in trap shown in (D, E, F) reveal binding and unbinding of motor. For each binding event, we determine the duration (ts ¼ t2  t1 Þ of the attached state, the mean load (F0), and the amplitude of load oscillations (ΔF) (See Sung et al. (2015) for details). (J, K, I) Data in (D, E, F) fitted with two harmonic functions, full line for attached state, dashed lines for unattached. Figure and legend are adapted from Sung, J., Nag, S., Mortensen, K. I., Vestergaard, C. L., Sutton, S., Ruppel, K. M., … Spudich, J. A. (2015). Harmonic force spectroscopy measures load-dependent kinetics of individual human β-cardiac myosin molecules. Nature Communications, 6, 7931. http://dx.doi.org/10.1038/ ncomms8931, with a Creative Commons license.

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Briefly, we used a 1064-nm fiber-coupled diode-pumped ND:YAG trapping laser beam (IPG Photonics, YLR-10–1064-LP), which was separated into two paths, one for each of the two traps, by a polarizing beam cube. An oil-immersion TIRF objective lens (Nikon, CFI Plan Apo 60) with high NA (NA ¼ 1.45) was used to tightly focus the trapping beams in the sample plane. Such beams can hold polystyrene beads near the focus with a Hookean spring-like force. We used an independent 845-nm fiber-coupled diode laser as detection beam (Lumics, LU0845) to measure the position of the beads via back-focal-plane (BFP) interferometry: a detection beam centered on a trapped bead produces an interference pattern at the BFP of the condenser. Each pattern is imaged on the sensor of a quadrant photodiode detector (QPD, Pacific Silicon Sensor, QP45-Q-HVSD). QPD outputs are in units of volt, which is why a calibration for volt-to-nm is required. The analog voltage signal was preamplified and antialiasing filtered (second order low-pass Butterworth) with a programmable filter (KROHN-HITE, 3944) with the cut-off frequency at the Nyquist frequency (20 kHz). The analog signal was digitized and sampled through a data acquisition card (National Instruments, PCIe-6363). Our normal sample rate of 40 kHz is sufficient to monitor fast dynamics and to perform trap calibration using powerspectral analysis. Bright-field illumination using a 740-nm LED (Mightex, LCS-0740-03-38) in conjunction with a CMOS camera (Thorlabs, DCC3240M) were used to image the beads in the sample chamber. Fluorescently labeled actin filaments were excited by a 532-nm diode laser and imaged with an EM-CCD camera (Andor, iXon Ultra 897). A highaccuracy, high-speed PZT (Physik Instrumente, P-545.3D7) was used to oscillate the sample chamber. We used a custom-built microscope body to mount the microscope components including the objective lens, condenser, PZT, LED, and QPD. Custom-written LabVIEW software was used to control most of the components and to display and record the data, including the positions of the beads and the stage. Schematics of the beam layouts and photos of the microscope are shown in Sung et al. (2010).

4. SAMPLE PREPARATIONS: PROTEINS, REAGENTS, AND BUFFERS 4.1 Human β-Cardiac Myosin S1 We expressed a short S1 construct (Sommese et al., 2013) of MYH7 (residues 1–808, truncated after the MYL3 binding site) followed by a 1  GSG flexible linker and an enhanced green fluorescent protein (eGFP) tag, which

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was coexpressed with MYL3 (ventricular essential light chain) with a FLAGtag for purification. Motors were further purified by a two-step purification procedure through actin sedimentation, the first with rigor attachment of motors to actin filaments without ATP and the second with inactive motors remaining bound to actin at high ATP concentration. Detailed protocols on cloning, virus production, and protein expression and purification are found elsewhere (Sommese et al., 2013; Sung et al., 2015).

4.2 Actin Filaments Actin was purified from fresh chicken-breast skeletal muscle, as described previously (Pardee & Spudich, 1982; Sommese et al., 2013). Purified actin was biotinylated to attach to NeutrAvidin-coated beads, and actin filaments were fluorescently labeled to be imaged under a fluorescence microscope. Surface exposed Cys residues on F-actin were covalently conjugated with biotin-maleimide, and subsequently labeled with TMR-Phalloidin. The detailed protocol is found in Sommese et al. (2013) and Sung et al. (2015).

4.3 Anti-GFP Antibody Anti-GFP antibody from Abcam (ab1218, 1 mg/mL or
6 μM) was used to attach the myosin-GFP specifically to the surface with the motor’s head oriented toward the actin dumbbell. The stock was aliquoted (2 μL), snapfrozen, and kept at 80°C. When to be used, one aliquot was thawed and diluted 100-fold (0.01 mg/mL or
60 nM) in assay buffer without BSA (see later), and was used directly for the trap experiments for a week or two without refreezing. We found that the binding specificity varies between different batches. We tested several different batches to find a satisfactory one and ordered from that (identified by its LOT number) from then on. Some batches showed a very high degree of nonspecific binding, which should be avoided.

4.4 Trapping Beads NeutrAvidin-coated polystyrene beads (1 μm in diameter) were used to trap and attach the biotin-labeled F-actin to form a dumbbell. We use commercially available NeutrAvidin-coated microspheres (Life Technologies, F8777), which have proven to work well for our trapping assay. These easily bind to biotin F-actin and we seldom observed aggregated beads.

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4.5 Nitrocellulose-Coated Coverslip A glass coverslip was first spin-coated with 1.5-μm diameter silica beads (Polysciences, 24327–15) after dilution to proper concentration in 0.1% Triton X-100. One such bead was used as a platform to support the motor in the three-bead assay. After the beads were dried on the surface, the coverslip was subsequently spin-coated again with 20 μL of nitrocellulose (EMS, 0.1% Collodion in amyl acetate). The nitrocellulose-coated glass coverslip was used to nonspecifically attach anti-GFP antibody to the surface, and subsequently it was blocked by BSA.

4.6 Other Reagents •

• • • • •

•

Milli-Q water or equivalent that has ultra-purity with high resistance (>14.3 MΩ). This water was used for buffers and reagents throughout the experiment. BSA: 10 mg/mL in water (10  stock). DTT: 1 M in water (100 stock). Unlabeled phalloidin: 100 mM in water (100 stock). ATP: 100 mM in water. pH was adjusted at (pH 7–7.5) using KOH. ATP regeneration system  Creatine phosphokinase (CPK): 10 mg/mL in AB with 50% glycerol (100  stock).  Phosphocreatine (PCR): 100 mM (25 mg/mL) in AB (100 stock). Oxygen-scavenging system  Glucose oxidase and catalase (GOC): 11 mg/mL glucose oxidase, and 1.8 mg/mL catalase in AB (100  stock).  Glucose: 20% (v/v) in water (100 stock).  Trolox: 25 mM in AB (20 stock)  Note that other oxygen-scavenging systems (e.g., PCA/PCD) can be used instead.

4.7 Assay Buffer (AB) Assay buffer (25 mM imidazole (pH 7.5), 25 mM KCl, 4 mM MgCl2, 1 mM EGTA, and 10 mM dithiothreitol (DTT)) was used for all trap experiments. We made a 10  AB stock without DTT and prepared fresh 1  AB with DTT prior to the experiments.

4.8 AB with BSA (ABBSA) AB with BSA (1 mg/mL) (ABBSA) is used to passivate the nitrocellulose surface after attaching the anti-GFP antibody to the surface. Surface

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blocking with BSA is to prevent any unwanted nonspecific attachment of other proteins including myosin and actin. We used a frozen aliquot of 10  BSA stock (10 mg/mL in water) to prepare fresh ABBSA before use.

4.9 GO buffer The final buffer for the trap experiments, termed GO buffer here, includes NeutrAvidin-coated polystyrene beads (104-fold diluted from the stock), TMR-phalloidin labeled biotinylated actin filaments (1–2 nM), nonfluorescent phalloidin (1 mM), ATP (2 mM) and an ATP regeneration system (0.1 mg/mL CPK and 1 mM PCR), and an oxygen-scavenging system (0.2% glucose, 0.11 mg/mL glucose oxidase, and 0.018 mg/mL catalase) in ABBSA.

5. EXPERIMENTAL PROTOCOLS The following procedures describe how to prepare and perform the experiment and collect data. For each step, we discuss important issues and how to trouble-shoot potential problems.

5.1 Preparation of Flow-Cell Sample Chamber The following steps prepare the sample chamber containing all the sample/ reagents: (1) Prepare buffers, samples, and reagents as described earlier. (2) Make a sample chamber with a glass slide and a nitrocellulose-coated coverslip attached with double-sided tape. The volume of the sample chamber is
10–15 μL. (3) Inject anti-GFP antibody (0.01 mg/mL in AB), which nonspecifically binds to the nitrocellulose surface. Incubate for 2 min. (4) Wash with ABBSA (5 volume) to passivate the sticky surface with BSA, which prevents nonspecific attachment of proteins except the preattached antibody. Incubate for 2 min. (5) Dilute the motor to an appropriate concentration. 100–500 nM (final concentration) is usually used for single molecule binding events. (6) Inject the motors through the chamber to let them bind to the antibody. Incubate for 2 min. (7) Wash with ABBSA (5 volume). (8) Inject GO buffer.

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(9) Seal the ends of the chamber with vacuum grease to avoid buffer evaporation during the experiment. (10) Mount the sample chamber firmly on the sample stage. (11) Image beads, actin in solution, and surface. • Image the beads and the actin filaments in solution to check their concentrations. If too high, then the beads or actin floating in bulk solution can interfere with the experiment. If too low, it becomes difficult to form a dumbbell. As a rule of thumb, one should observe only a few beads and filaments in each field of view. • Check whether many filaments are stuck to the surface, as this may indicate the presence of inactive motors or nonspecific sticking to the anti-GFP antibody. • Check the density of platform beads on the surface. Fast spin coating helps to spread the beads uniformly and prevents bead aggregation as the solution dries out. • Surface passivation is a critical step and is context dependent. Surface and filament interaction depends on many factors, including the electrical charge, hydrophobicity, pH, and salt condition. Surface passivation with BSA has been commonly used in the field of actin-based myosin motility. Other surface passivation can be used alternatively, e.g. PEG, pluronic acid, or casein, depending on the types of protein and the filament. For example, casein has been commonly used in microtubule-based kinesin or dynein motility, and BSA does not work in this system (personal communication).

5.2 Formation of an Actin Dumbbell The following steps lead to a dual-trap and a stable actin dumbbell for HFS. The experiment is described in the next subsection. (1) Trap two beads near the surface, one bead in each trap. • Adjust the power of the trapping beam. If too weak, it cannot trap a bead. If too strong, it can cause damage to the optics or electronics in the system. Higher power, though, increases stability. A typical range of trap power in our experiment was 100–200 mW when measured at a position right before the objective lens. • Trapping is possible only near the surface of the glass coverslip, within a few micrometer where aberration due to the index mismatch at the interface between glass and water is not significant for the employed oil-immersion objective lens. Since the HFS

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experiment is done near the surface, this is not a problem. A water immersion lens would allow trapping deep in solution. • It is only possible to trap beads below the focal plane since the radiation force pushes the beads in the direction of the beam propagation, which is upward in our inverted microscope system. In our imaging system, a bead below the image plane looks dark in the central area of its image, while beads above the plane look bright in the same central area. • If the beam alignment is done properly, then the focal plane of the trap and the detection beams are near the sample plane of the imaging system. The trapped bead should remain in focus. Otherwise, adjust the focal depth of the trap/detection beam and the imaging system. (2) Separate the beads by 3–5 μm, appropriate to form an actin dumbbell in the next step. • If too close, then the two trapped beads might interfere with each other as well as with the platform bead on the surface. • If too far apart, then it becomes difficult to find a long filament to bridge them and the compliance in the dumbbell becomes significant. (3) Position the detection beams to the center of the trapped beads. • Move the detection beam in (x, y) direction using the piezo mirror in the detection beam path and find the center of the detection signal. • When the trap and the detection beams are coaligned, choose a pair of trap and detection beams with orthogonal polarization (p or s) to minimize the risk of interference between the trap and the detection beams. • If the trap and the detection beam alignment is done properly and the trapped bead is spherically isotropic, then the (x, y) detection signal should show effectively a sinusoidal-like signal in the overall range and a linear signal near the center. If not, then release and trap another bead to check the possibility of a bead-specific problem (nonspherical bead or junk stuck to the bead), multiple beads trapped, or a bubble near the laser focus. If the problem is systematic, then realign the trap and the detection beam paths. • Detection beam power should be strong enough to get highresolution and low-noise signals, without affecting the trapping and interfering with it.

Single-Molecule Harmonic Force Spectroscopy

•

15

Check the Brownian fluctuations of the trapped beads. If the signal looks noisy, especially in the low frequency range, junk might be stuck to the beads. Real-time display of the power spectrum helps to identify the noise level. (4) At this stage, we normally calibrate the force from each trap, due to variable bead properties. We usually do this calibration before taking data with myosin in case the beads are lost over the course of the experiment. The calibration protocol is described in the next section. After calibration, we form an actin dumbbell. (5) Move the sample stage to attach biotin-labeled actin diffusing in solution to the trapped NeutrAvidin-coated beads. Attach each end of the filament to each bead to form an actin dumbbell. • We do bright-field imaging of beads and surface via LED and a CMOS camera and fluorescence imaging of actin via a 532-nm diode laser and an EM-CCD camera. Visualize them together to check coordinated positions of beads and actin. • To form a dumbbell, first, find an actin filament with an appropriate length (3–5 μm). Place a bead near one end of the filament. Sometimes it is time consuming and difficult to make the attachment as the filament moves around and easily diffuses away from the focus. Be patient and move on to the next filament until catching one. If one end of a filament is attached, then move the stage in the opposite direction at a constant speed, such that the free end of the filament comes close to another bead. Once in a while move the stage in the orthogonal direction of the dumbbell to check the double attachment. • Sometimes, beads bind around the middle of the filament. In this case, it is difficult to form a dumbbell, and it is preferable to release the bead and trap a new one. • Be careful to avoid accidental trapping of additional free beads in solution. It causes release of the trapped bead or aggregation of the two beads. • Radiation pressure from the trap often pushes the actin filament into the solution, away from the trapped beads. If this continues, then temporarily reduce the trapping power until the actin filament is attached. • Moving deep in solution, a few micrometer further, increases the chance of finding more actin filaments. The trapping force, however, becomes weak due to aberrations. Be careful not to move too deep.

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•

Abrupt and fast stage movement can cause escape of the beads from the trap due to the drag force from the fluid. Move beads at an appropriate speed and acceleration to avoid their accidental loss. • Sometimes actin does not stick to the beads. If biotin or NeutrAvidin is not fresh, then prepare fresh samples. Often, the biotin-labeling efficiency on actin is not high and there is not enough biotin on the filament. Prepare new biotin-actin with fresh biotin-maleimide. Test the biotin-labeling efficiency by attaching the filament to a StreptAvidin or NeutrAvidin-coated surface on the coverslip. • Check the time traces of bead positions and their power spectra. If the position signal is much noisier than previously, then a short filament or junk may have stuck to the bead. Trap new beads in this case. • Forming a dumbbell in a timely manner (ideally in 10–20 min) is critically important to increase the chance of success for collecting quality data. Human β-cardiac myosin stays active for an hour or two at room temperature. After that, a new sample should be prepared. (6) Make a tightly stretched dumbbell by separating the two beads. • To make a tight dumbbell, first, oscillate one of the beads (call it Bead 1) via the AOD. The amplitude of this oscillation should be two or three times the amplitude of the bead’s Brownian motion in the trap. If the dumbbell is not stretched, the other bead (Bead 2) does not oscillate in response. Second, in that case, use the piezo mirror to move Bead 1 slowly to increase the interbead distance. Third, move Detection Beam 1 to the center of Bead 1. Fourth, repeat the second and the third steps until Bead 2 oscillates when Bead 1 is oscillated. Finally, carefully move Bead 1 until the oscillation signals of the two beads are similar. In the ideal limit of a completely stretched dumbbell with no compliance, bead-to-bead correlation is complete. • Be very careful not to break the actin. Actin becomes fragile under strong tension when it is fully stretched. If this happens, trap new beads and make a new dumbbell. • Sometimes only one of the beads is lost while the actin dumbbell is still connected. This happens frequently when the stage suddenly moves while the dumbbell is fully stretched. Retrapping the straying bead is, however, not easy due to the radiation force pushing on it during attempts. If one dumbbell bead goes astray, first turn off the

Single-Molecule Harmonic Force Spectroscopy

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trapping beam of the lost bead. Then, reduce the distance between the two traps. Move the stage in the direction of the empty trap, such that the fluid drag force on the dumbbell drags it toward capture in both traps. Then, suddenly turn on the beam of the empty trap again. This will allow the retrapping of the lost bead while maintaining the dumbbell. Since the dumbbell is relaxed, it needs to be stretched again as described ealier.

5.3 HFS: Experiment The HFS experiment roughly consists of three steps: searching for an active motor, executing HFS, and data collection. (1) Move the dumbbell close to a platform bead and find an active motor. • To find an active motor, first, slowly move the stage such that a platform bead is placed between the two trapped beads of the dumbbell. Second, lower the dumbbell close to the platform bead’s surface such that a motor attached to the platform bead can bind to the actin filament. Third, sinusoidally oscillate the PZT, which amplifies the binding signal to allow easy detection of the motor binding event. It is important to position the dumbbell at an appropriate height not too close to the surface, since that may cause signals from nonspecific sticking. If no binding is observed, stop the oscillation, move it to the next platform bead, and oscillate the PZT again. Repeat these steps until a binding event that appears to be from an active motor is observed. Finally, stop the oscillation and confirm that it is indeed from an active motor. The latter produces a unidirectional stroking signal, while nonspecific sticking does not. Once an active motor has been confirmed, one is ready to execute the experiment and collect data. • Be careful to avoid signals from nonspecific sticking since they contaminate the actual signal. A too-short distance between dumbbell and platform beads, an elevated amplitude of oscillation, and an elevated frequency of attachment all indicate nonspecific sticking. Certain batches of anti-GFP antibody seem to cause sticking with a higher probability, as does higher concentration of the antiGFP antibody. We tried to avoid false-positive signals with these observations in mind. • Before doing the trapping experiment, we usually start with the motility assay (actin gliding driven by motors attached to the

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surface). Observation of smooth actin gliding in the motility assay is a prerequisite for the success of the single-molecule experiment, as our three-bead assay with a dual-beam actin dumbbell is simply a single-molecule version of the motility assay. • We do multiple control experiments to make sure that the signal is from an active motor. Parameters to control include motor and ATP concentrations. Start with high concentration of motor at low ATP concentration, since this shows more frequent and longer binding, easy to detect even without the PZT oscillation. Then, lower the motor concentration at low ATP concentration to find the optimal motor concentration. As a rule of thumb, one should find one motor binding among 10–20 platform beads. Finally, use the same condition and gradually increase the ATP concentration till saturated (2 mM), which results in short binding time. (2) Execute the HFS experiment with the single active motor. • Single-molecule HFS is simply a continuation of the myosin–actin binding events with PZT oscillation, performed in a more controlled manner. Once an active motor is confirmed, we repeat the sinusoidal oscillation of the PZT at an appropriate amplitude and frequency, and record data until the motor becomes inactive. We oscillated the PZT with an amplitude of 30–50 nm at a frequency of 100–200 Hz in our previous study. While strongly bound, myosin is under an oscillation force with a mean force that depends on where in the oscillation cycle the binding happened. This way, randomly selected mean forces are applied in every binding event. The amplified signal of the bead position with the harmonic oscillation of the PZT makes it easy to detect the binding and measure the dwell time accurately with an automatic binding detection algorithm (see next section). • It is important to maintain the dumbbell position at the appropriate position in 3D, where the single motor has binding access to the actin without spatial constraints. The dumbbell should not be too close to the platform bead: so close that the signal becomes noisier and nonspecific sticking might happen. At an appropriate distance and position, motor binding happens frequently, e.g., once every second on average. Optimal statistics are achieved with the highest frequency of true binding for which one can distinguish consecutive binding events. • Drift of the sample chamber can occur over the course of the data collection. If the binding frequency drops, then stop data collection,

Single-Molecule Harmonic Force Spectroscopy

•

•

•

19

fine tune the position in 3D using the PZT, and recollect data, if the binding signal is recovered with the adjustment. It is important to know the actual amplitude of the PZT oscillations. Our PZT has a resonance frequency at 500 Hz in (x, y) according to its specification, and its amplitude of oscillation is attenuated as the frequency of oscillation of the voltage driving the stage approaches the resonance frequency. This is due to a built-in low-pass filter, a safeguard against potential damage to the PZT. Consequently, the amplitude specified for a given voltage may differ in a frequencydependent manner from the actual amplitude achieved. Please note that the resonance frequency value depends on the load applied to the PZT, so the information in the specification might be different from the actual value. Our PZT outputs a sensor signal in volt as a read-out of the position of the PZT. The volt-to-nm conversion factor of the sensor signal was precalibrated by the manufacturer (Physik Instrumente) and we confirmed it, using a calibration bar. We then characterized the frequency-dependent amplitude changes in our system. For example, when we oscillated the PZT at 200 Hz with amplitude 50 nm, the actual oscillation amplitude was around 35 nm. We used the latter value in our study. It is also important to carry out the experiment at different oscillation amplitudes and frequencies to find a regime in which it works. For example, if the amplitude of oscillations is too large, then the force applied to the motor exceeds the motor’s stall force, which can result in premature detachment from the actin. Also, our theory assumes that the motor is in a quasi-steady state under the applied oscillations at frequencies of 100–200 Hz, which means that the motor continuously is in a configuration adapted to the instantaneous load. This proved to be true in our experiment. However, this assumption might not be satisfied at much higher frequencies. We used a PZT to apply a harmonic oscillation, but one could use other methods. In fact, an AOD or AOM can also be used to oscillate the dumbbell directly with increased oscillation frequency and temporal resolution, which would be useful to measure the loaddependence of even shorter and weaker events.

6. TRAP CALIBRATION We recorded data from the detector in volts, which needs to be converted to nanometers. Each trap beam pulls a bead similarly to a

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Hookean spring, so one needs to determine the spring constant. Calibration of the trap is critical in order to obtain accurate information of the force in pico-newtons (pN) and displacement in nanometers (nm). We obtain two pieces of information from the following calibration procedure, one is the volt-to-nm conversion factor for the detector (QPD) and the other is the trap stiffness (or spring constant) in pN/nm. (1) After trapping a bead in each trap, we move it close to the surface where the HFS experiment is usually carried out. Both the trap stiffness and the volt-to-nm conversion factor are functions of distance from the surface, so it is important to calibrate at the right height. For example, the trap stiffness grows weaker with distance from the surface due to aberrations in our microscope system. (2) We calibrate the QPD by raster scanning of each bead in 2D using the AOD (or AOM), e.g., we displace the beads by 60-nm steps over an area of 600  600 nm. Meanwhile, we simultaneously record both the QPD output signal (x and y in volt) and the bright-field images of the beads (in pixels). Centroid tracking of the bead displacement (in pixels) and imaging a calibration bar (pixels to nanometers) allowed us to convert the bead displacement from pixels to nanometers. We then correlated the linear relationship between the QPD signal (in volt) and the bead displacement (in nanometers) to obtain the volt-to-nm conversion factor. (3) We estimate the trap stiffness by maximum-likelihood fitting of the theoretical power spectrum to the measured power spectrum of the recorded thermal motion of the beads in the traps (Nørrelykke & Flyvbjerg, 2010; Toli
c-Nørrelykke et al., 2006). We used the QPDs to record the positions of the beads for
25 s (106 data points at 40 kHz sampling frequency) and calculated the power spectra of the recorded time series using windowing (4000 data points in each window) as described in Section 12.7 of Press, Teukolsky, Vetterling, and Flannery (1986). We fitted each power spectrum with a sum of two Lorentzians while taking into account low-pass filtering by our antialiasing filter and aliasing of the filtered signal (Berg-Sørensen & Flyvbjerg, 2004). One Lorentzian models low frequency noise in the setup due to air fluctuation and vibration. The other Lorentzian extracts the trap stiffness and the bead’s drag coefficient near the surface (Sch€affer, Nørrelykke, & Howard, 2007; Toli
c-Nørrelykke et al., 2006).

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7. HFS: THEORY AND DATA ANALYSIS The next step is to extract the force-dependent kinetic information from the recorded data. The original data are time traces of the trapped beads’ positions in volts. With the calibration information, we can convert the bead positions to nanometers by multiplication with the QPD factor in nm/V, and also measure the force applied to the motor in pico-newtons by multiplication of the trap stiffness in pN/nm. The time trace displays multiple independent myosin binding events with large amplitude harmonic oscillations, separated by unbound states with smaller oscillation amplitude and different phase in response to the drag force from the buffer’s oscillatory motion with the sample chamber (Fig. 3D–F). Force–kinetics relationships can be investigated by plotting the bound time (inverse kinetic rate) vs the mean force applied during that time. To that end, we need good statistics, i.e., we need to detect a large number of binding events effectively and accurately.

7.1 Automatic Binding Detection We detect events of myosin binding to the actin dumbbell by monitoring the displacements of the individual bead in its optical trap (Fig. 3D–F). When the myosin is not bound to the dumbbell, the dumbbell oscillates due to drag from the fluid in the sample chamber (Fig. 3D–F, left and right portions of the traces), which follows the stage motion. In this oscillation, the dumbbell position follows the fluid velocity closely and is therefore ahead of the stage position by almost π/2, see Eq. (7) in Toli
c-Nørrelykke et al., 2006. When the myosin is bound to the actin dumbbell, the oscillation of the PZT is transferred to the trapped beads, and the positions of the beads are now almost in phase with the oscillating position of the stage (Fig. 3D–F, center portions of the traces)—compliance of the dumbbell allows the beads to get a little ahead of the stage position, dragged by the fluid in the sample chamber, but the net effects of binding are oscillations of the dumbbell with much larger amplitude, typically around a different mean position, and a phase shift by almost π/2 relative to the phase of oscillations in the unbound state (Fig. 3G–I, black traces). Thus, we have two criteria for selection of binding events: changes in phase (Fig. 3G–I, black traces) and changes in amplitude (Fig. 3G–I, alternatively colored traces), like the signals of AM and FM radio, respectively, except we “listen” to both simultaneously.

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We developed an automatic binding detection algorithm that finds true motor binding events in the presence of noise from full time traces. This method is superior to manual selection since it is more objective, efficient, and accurate. The software that executes this algorithm is freely available via http://www.nanotech.dtu.dk/Research-mega/Forskningsgrupper/SSS. Details about its signal processing are given in the methods section of Sung et al. (2015).

7.2 Force-Dependent Kinetics Under Harmonic Force HFS hinges on two points: (i) Because of the vast separation of scales between the myosin molecule and the microscope stage, our harmonic motion of the stage is extremely slow as experienced by the motor. So while we vary the force exerted on the motor harmonically, the motor experiences a quasi-stationary situation. The force appears as if clamped on the motor’s time scale—for long enough, anyways, for the motor to be in equilibrium with the instantaneous force at every instant. Consequently, the rate of detachment of the motor from actin is, at each instant, the rate we would observe, if the force were clamped with the value it has at that instant. This is an assumption, but a plausible one, as just argued, and it is tested experimentally when we use it. (ii) This assumption makes force-clamping superfluous. It is replaced with a little bookkeeping of rates that vary with varying loads. This goes on continuously in time, so a differential equation appears. Our second point is that we can handle this, and we do so in the next subsection. Force-clamping reduces this bit of math to elementary arithmetic. But one should not complicate experimental design just to simplify the math of one’s data analysis, because math is pure logic. It processes data without adding noise, which makes it better than most experimental tools. Moreover, in the specific case at hand, the assumed quasi-stationarity simplifies kinetics to such an extent that the release rate for actin from myosin when experiencing a harmonic load is the same function as for a constant load, except the mean load replaces the constant load, and an overall correction factor describes the increase in release rate resulting from harmonic oscillations in load; see next subsection.

7.3 Mathematical Model and Theory The mean value F0 of the oscillating load on the myosin depends on where in the cycle of the periodic motion of the stage the myosin happens to attach to the dumbbell. F0 equals minus the sum of the mean forces from the two

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traps on the dumbbell, which we measure. Thus, we measure the lifetime of the attached state of myosin for a range of measured F0-values. Fig. 4A shows a scatter plot of this F0 vs lifetime. It is interpreted as follows: Let k(F) denote the load-dependent rate of unbinding of myosin from actin. Let F be a function of time t. Then the “survival function” P(t1, t2) of the attached state— i.e., the probability of remaining attached from time t1 of binding till a later time t2—evolves in time according to the quasi-stationary dynamics d P ðt1 , tÞ ¼ kðF ðt ÞÞP ðt1 , t Þ: dt This differential equation is solved by  Z t2  kðF ðt ÞÞdt : P ðt1 , t2 Þ ¼ exp 

(1)

(2)

t1

The probability density function (pdf ) on the time axis for unbinding at a time t, given binding took place at time t1, is equal to minus the right-hand side in Eq. (1). At constant load F, as in force-clamp spectroscopy, this pdf is a simple exponential function of the difference t2  t1. For time-dependent F, this pdf is the function that results from inserting Eq. (2) in the right-hand side of Eq. (1). If F is a periodic function of t, and t2  t1 ¼ ntdrive is an integer number of periods tdrive of F, then 



P ðt1 , t2 Þ ¼ ekn tdrive ¼ ek ðt2 t1 Þ ,

(3)

where k denotes the time average of k(F(t)) over a period. This is a simple exponential function of n. In our data analysis, we use the probability of unbinding in the period following n periods in the bound state, which is 

P ðt1 , t2 Þ  P ðt1 ,t2 + tdrive Þ ¼ C ekntdrive ,

(4)



where C ¼ 1  ektdrive . This also is a simple exponential function of n, known as the geometric distribution. It shows that “periodic force spectroscopy,” as we call it, in its data analysis is as simple as force-clamp spectroscopy. Its use only requires that the period, tdrive, is shorter than the timescales one wishes to resolve. We achieve further simplifications by using a periodic load that is harmonic, in what we call “Harmonic Force Spectroscopy,” F ðtÞ ¼ F0 + ΔF sin ð2πfdrive ðt  tM ÞÞ:

(5)

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It has fdrive ¼ 1=tdrive and tM denoting the arbitrary phase of the oscillating stage. As ΔF differs little between binding events, F0 is the only quantity in this expression that really differs between binding events. Eq. (4) therefore predicts that binding events which have (nearly) the same mean load F0, have attachment times that are exponentially distributed in time, when binned on the time axis in bins that last an integer number of periods of the stage motion, e.g., one period. Fig. 4B shows that this is, indeed, the case. In this manner, we determine k experimentally for a range of values for F0. Fig. 4C shows a plot of these values. This plot describes a loaddependent ADP release rate that is convincingly described by an exponential dependence on the load F0. This observation points toward Arrhenius’ equation,   Ea k ¼ A exp  kB T

(6)

and transition state theory. In Eq. (6), Ea is the activation energy, the energy difference between the bound state and the transition state for detachment. If a constant external load F opposes the transition, Ea is increased by the amount of work done by the myosin against that load. With δ denoting the distance between the bound state and the transition state, as measured along the reaction pathway, the height of the energy barrier toward unbinding is increased from Ea to Ea + Fδ. Consequently,   Fδ kðF Þ ¼ k0 exp  kB T

(7)

where k0 is the rate at zero load, and hence, with our harmonic force in Eq. (5),     ΔFδ F0 δ  k ¼ kðF0 , ΔF Þ ¼ k0  I0 exp  kB T kB T

(8)

where I0 is the zeroth-order modified Bessel function of the first kind. Thus, we see that Arrhenius’ equation explains the observed exponential dependence of k on the mean load F0. Since kBT and ΔF are known, a fit of Eq. (8) to data, using k0 and δ as fitting parameters, determines these two parameters of the load-dependent ADP release rate; see Fig. 4C.

A

B

C

Fig. 4 Load-dependent ADP release rate measured with 200-Hz stage oscillation. (A) Scatter plot of duration of binding (ts) vs load (F0) (N ¼ 388). (B) Histogram of durations ts in the narrow range of loads in the red box in panel (A) (N ¼ 41). Histogram bins are 5 ms wide, i.e., one period of the stage motion. The histogram is fitted by Eq. (4) using maximum-likelihood estimation (MLE) to obtain k ¼ k ðhF0 i, ΔF Þ. The histogram is consistent with an exponential distribution (full line). The characteristic time of this exponential is the inverse of the ADP release rate under the average load in the red box in panel (A),  k ¼ k ðhF0 i, ΔF Þ. (C) ADP release rates depend exponentially on applied load. Black data points show ADP release rates k ¼ k ðhF0 i, ΔF Þ against mean load hF0i for each of 10 consecutive 1-pN bins. The individual error bars are calculated from the variance of the MLE in (B) as the inverse Fisher information for the parameter (Sung et al., 2015). The full curve is the fit of Eq. (8) to the rates, yielding k0 ¼ 71  4s1 , δ ¼ 1:01  0:09nm, corresponding to k ð0, ΔF Þ ¼ 89s1 on the full curve. Figure and legend are adapted from Sung, J., Nag, S., Mortensen, K. I., Vestergaard, C. L., Sutton, S., Ruppel, K. M., … Spudich, J. A. (2015). Harmonic force spectroscopy measures load-dependent kinetics of individual human β-cardiac myosin molecules. Nature Communications, 6, 7931. http://dx.doi.org/10.1038/ncomms8931, with a Creative Commons license.

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8. RESULTS AND DISCUSSION We found that the force-dependent ADP release rate of human β-cardiac S1 fits well to a single exponential function (Fig. 4C), consistent with the transition state theory and the Bell–Evans model for bond rupture. A fit of Eq. (8) to the data yielded k0 ¼ 71  4s1 and δ ¼ 1:01  0:09nm (Fig. 4C). We obtained the fitting results from seven independent measurements with six different molecules (Fig. 5A and B). They average to A

B

Fig. 5 Individual parameter values for seven independent measurements with six myosin molecules. Each molecule is analyzed as in Fig. 4. The stage is oscillated at 200 Hz except for #1 (100 Hz). The number of attachment events for each molecule are, respectively, N ¼ 138, 388, 569, 174, 539, 950, and 229. Figs. 3 and 4 use data for Molecule #2. Points with error bars denote single-molecule results. The error bars are obtained from the theoretical covariance matrix (¼inverse Fisher information matrix) of the singlemolecule estimates for k0 and δ (Sung et al., 2015). (A) The load-free ADP release rates k0, corrected by I0 ðΔFδ=kB T Þ to adjust for oscillation effects. Their weighted mean value, k0 ¼ 87  7s1 , is shown as a horizontal full line with s.e.m. shown as a shaded area. If Molecule #6 is treated as an outlier and dropped from our statistics, we find k0 ¼ 80  7s1 . The unloaded ADP release rate measured in stopped-flow experiments, k0 ¼ 72  5s1 , is shown as a horizontal dashed line with s.e.m. shown as a shaded area. (B) The distance δ from the strongly bound state of each myosin to its transition state toward ADP release, measured along the reaction path. Their weighted mean value, δ ¼ 0:8  0:1nm, is shown as a horizontal full line with s.e.m. shown as a shaded area. If Molecule #6 is treated as an outlier and dropped from our statistics, we find δ ¼ 0:7  0:1nm. Figure and legend adapted from Sung, J., Nag, S., Mortensen, K. I., Vestergaard, C. L., Sutton, S., Ruppel, K. M., … Spudich, J. A. (2015). Harmonic force spectroscopy measures load-dependent kinetics of individual human β-cardiac myosin molecules. Nature Communications, 6, 7931. http://dx.doi.org/10.1038/ncomms8931, with a Creative Commons license.

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k0 ¼ 87  7s1 (Fig. 5A, blue) and δ ¼ 0:8  0:1nm (Fig. 5B) (mean s.e.m., N ¼ 7). The unloaded ADP release rate also agrees with our stopped-flow experiment, done in solution and hence unloaded (Fig. 5A). Our protocol enabled recording of a sufficient number of attachment events for individual molecules—between 138 and 950—to allow this statistical analysis for individual molecules. This is an essential improvement of a single-molecule experiment: this step from ensemble results to singlemolecule results allows us to inspect the performance of individuals in an ensemble before we average experimental results over the ensemble to improve statistics. By doing that, statistical outliers can be eliminated from the ensemble before averaging, which may improve accuracy and precision of ensemble-averaged results dramatically, depending on the frequency and extremity of outliers. Molecule #6 in Fig. 5 provides an example: its elimination as an outlier shifts the ensemble-averaged value for k0 by one standard error.

9. CONCLUSION AND OUTLOOK HFS has proven to be useful for obtaining the load-dependent rate at the single-molecule level in an easy, fast, and efficient way. The method was applied to demonstrate the load-dependent ADP release rate of human β-cardiac S1. The exponential change of rate provides a clue to the molecular basis of the force–velocity curve and the Fenn effect of contracting muscle. In a translational point-of-view, the method can be applied to human β-cardiac S1 bearing hypertrophic cardiomyopathy (HCM)-causing singlepoint mutations to understand the devastating heart disease at the singlemolecule level (Spudich, 2014; Spudich et al., 2016). This method is not limited to our myosin study but can be further extended into other systems, with some modification and optimization. In this method chapter, we aimed to provide a detailed protocol for how to carry out the experiment. We hope that this is useful to others who want to apply HFS to their own experimental system.

ACKNOWLEDGMENTS We thank Suman Nag, Christian L. Vestergaard, Shirley Sutton, and Kathleen Ruppel for assisting the experiment, the data analysis, and the sample preparation. This work was funded by the National Institutes of Health (NIH) Grant R01 GM033289 and NIH Grant R01 HL1171138 (to J.A.S.), the Human Frontier Science Program GP0054/2009-C (to J.A.S. and H.F.), a Stanford Bio-X Fellowship (to J.S.), and a Lundbeck Fellowship (to K.I.M.).

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REFERENCES Bell, G. I. (1978). Models for the specific adhesion of cells to cells. Science, 200(4342), 618–627. Berg-Sørensen, K., & Flyvbjerg, H. (2004). Power spectrum analysis for optical tweezers. Review of Scientific Instruments, 75(3), 594–612. Capitanio, M., Canepari, M., Maffei, M., Beneventi, D., Monico, C., Vanzi, F., … Pavone, F. S. (2012). Ultrafast force-clamp spectroscopy of single molecules reveals load dependence of myosin working stroke. Nature Methods, 9(10), 1013–1019. Capitanio, M., & Pavone, F. S. (2013). Interrogating biology with force: Single molecule high-resolution measurements with optical tweezers. Biophysical Journal, 105(6), 1293–1303. http://dx.doi.org/10.1016/j.bpj.2013.08.007. Finer, J. T., Simmons, R. M., & Spudich, J. A. (1994). Single myosin molecule mechanics: Piconewton forces and nanometre steps. Nature, 368, 113–119. Greenberg, M. J., Shuman, H., & Ostap, E. M. (2014). Inherent force-dependent properties of ß-cardiac myosin contribute to the force-velocity relationship of cardiac muscle. Biophysical Journal, 107(12), L41–L44. http://dx.doi.org/10.1016/j.bpj.2014.11.005. Greenleaf, W. J., Woodside, M. T., & Block, S. M. (2007). High-resolution, single-molecule measurements of biomolecular motion. Annual Review of Biophysics and Biomolecular Structure, 36, 171–190. http://dx.doi.org/10.1146/annurev.biophys.36.101106.101451. Neuman, K. C., & Block, S. M. (2004). Optical trapping. The Review of Scientific Instruments, 75(9), 2787–2809. http://dx.doi.org/10.1063/1.1785844. Neuman, K. C., & Nagy, A. (2008). Single-molecule force spectroscopy: Optical tweezers, magnetic tweezers and atomic force microscopy. Nature Methods, 5(6), 491–505. http:// dx.doi.org/10.1038/NMETH.1218. Nørrelykke, S. F., & Flyvbjerg, H. (2010). Power spectrum analysis with least-squares fitting: Amplitude bias and its elimination, with application to optical tweezers and atomic force microscope cantilevers. The Review of Scientific Instruments, 81(7), 075103. http://dx.doi. org/10.1063/1.3455217. Pardee, J. D., & Spudich, J. A. (1982). Purification of muscle actin. Methods in Enzymology, 85, 164. Press, W. H., Teukolsky, S. A., Vetterling, W. E., & Flannery, B. P. (1986). Numerical recipes. The art of scientific computing. Cambridge: Cambridge University Press. Sch€affer, E., Nørrelykke, S. F., & Howard, J. (2007). Surface forces and drag coefficients of microspheres near a plane surface measured with optical tweezers. Langmuir: The ACS Journal of Surfaces and Colloids, 23(7), 3654–3665. http://dx.doi.org/10.1021/la0622368. Sommese, R. F., Sung, J., Nag, S., Sutton, S., Deacon, J. C., Choe, E., … Spudich, J. A. (2013). Molecular consequences of the R453C hypertrophic cardiomyopathy mutation on human β-cardiac myosin motor function. Proceedings of the National Academy of Sciences of the United States of America, 110(31), 12607–12612. http://dx.doi.org/10.1073/ pnas.1309493110. Spudich, J. A. (2014). Hypertrophic and dilated cardiomyopathy: Four decades of basic research on muscle lead to potential therapeutic approaches to these devastating genetic diseases. Biophysical Journal, 106(6), 1236–1249. http://dx.doi.org/10.1016/j.bpj.2014.02.011. Spudich, J. A., Aksel, T., Bartholomew, S. R., Nag, S., Kawana, M., Yu, E. C., … Ruppel, K. M. (2016). Effects of hypertrophic and dilated cardiomyopathy mutations on power output by human β-cardiac myosin. Journal of Experimental Biology, 219(2), 161–167. http://dx.doi.org/10.1242/jeb.125930. Sung, J., Nag, S., Mortensen, K. I., Vestergaard, C. L., Sutton, S., Ruppel, K. M., … Spudich, J. A. (2015). Harmonic force spectroscopy measures load-dependent kinetics of individual human β-cardiac myosin molecules. Nature Communications, 6, 7931. http://dx.doi.org/10.1038/ncomms8931.

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Sung, J., Sivaramakrishnan, S., Dunn, A. R., & Spudich, J. A. (2010). Single-molecule dualbeam optical trap analysis of protein structure and function. Methods in Enzymology, 475(10), 321–375. http://dx.doi.org/10.1016/S0076-6879(10)75014-X. Toli
c-Nørrelykke, S. F., Sch€affer, E., Howard, J., Pavone, F. S., J€ ulicher,, F., & Flyvbjerg, H. (2006). Calibration of optical tweezers with positional detection in the back focal plane. Review of Scientific Instruments, 77(10), 103101. Veigel, C., Molloy, J. E., Schmitz, S., & Kendrick-Jones, J. (2003). Load-dependent kinetics of force production by smooth muscle myosin measured with optical tweezers. Nature Cell Biology, 5(11), 980–986. http://dx.doi.org/10.1038/ncb1060.

CHAPTER TWO

Studying the Mechanochemistry of Processive Cytoskeletal Motors With an Optical Trap V. Belyy*, A. Yildiz†,1 *Biophysics Graduate Group, University of California at Berkeley, Berkeley, CA, United States † University of California at Berkeley, Berkeley, CA, United States 1 Corresponding author: e-mail address: [email protected]

Contents 1. Introduction 1.1 Cytoskeletal Motors 1.2 Using an Optical Trap for Single-Motor Measurements 1.3 Measuring Motor Stall Forces 1.4 Force–Velocity Measurements Using Force Feedback 1.5 Studying Interhead Coordination Using Force-Dependent Unbinding Data 2. Experimental Setup and Troubleshooting 2.1 Optical Layout of the Trap 2.2 Software Control of the Trap 2.3 Noise Elimination 3. Experimental Protocols 3.1 Coating Polystyrene Beads With Antibodies 3.2 Human Dynein Optical Trapping Sample Preparation 3.3 Trap Position vs AOD Command Signal Calibration 3.4 Trap Stiffness and PSD Response Calibration 3.5 Dynein Stall Force Measurements 3.6 Identification of Motor Stalls 3.7 Force-Feedback Assay 3.8 Construction of Force–Velocity Curve 3.9 Square Wave Unbinding Force Assay 3.10 Analysis of Unbinding Force Data 4. Conclusion Acknowledgments References

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Abstract Cytoskeletal motors utilize the energy stored in ATP to generate linear motion along rigid filaments. Because their enzymatic cycles are tightly coupled to the production of force and forward movement, the optical-trapping technique is uniquely suited for studying their mechanochemical cycle. Here, we discuss the practical aspects of optical trapping in connection with single-motor assays and describe three distinct experimental modes (fixed-trap, force feedback, and square wave) that are typically used to investigate the enzymatic and biophysical properties of cytoskeletal motors. The principal outstanding questions in the field involve motor regulation by cargo adaptor proteins and cargo transport by teams of motors, ensuring that the optical trap’s ability to apply precise forces and measure nanometer-scale displacements will remain crucial to the study of intracellular motility in the foreseeable future.

1. INTRODUCTION 1.1 Cytoskeletal Motors Processive cytoskeletal motors are a unique class of enzymes whose primary output is mechanical work performed by pulling a cargo along a rigid linear track. As a result, the study of these motors has greatly benefitted from biophysical methods that enable both measurement and application of precise forces at the single-molecule level. Optical trapping remains the most versatile and widely used method with such capabilities. It has been successfully used to interrogate the mechanochemical cycles of motors from the kinesin (Visscher, Schnitzer, & Block, 1999), dynein (Gennerich, Carter, ReckPeterson, & Vale, 2007), and myosin (Clemen et al., 2005) families. We focus on cytoplasmic dynein in this chapter, but the general principles were successfully applied to other motor proteins in the past two decades. Cytoplasmic dynein (simply “dynein” hereafter) is a dimeric protein that harnesses the energy of ATP hydrolysis to walk along microtubules in the minus-end direction and pull cellular cargoes with forces ranging from
1 to
4 pN (Belyy, Hendel, Chien, & Yildiz, 2014; Belyy et al., 2016; McKenney, Vershinin, Kunwar, Vallee, & Gross, 2010; Ori-McKenney, Xu, Gross, & Vallee, 2010; Rai, Rai, Ramaiya, Jha, & Mallik, 2013; Torisawa et al., 2014), depending on the presence of regulatory proteins such as dynactin and BICD2N (McKenney, Huynh, Tanenbaum, Bhabha, & Vale, 2014; Schlager, Hoang, Urnavicius, Bullock, & Carter, 2014). Unlike kinesin-1 (Yildiz, Tomishige, Vale, & Selvin, 2004) and myosin V motors (Yildiz et al., 2003) the enzymatic cycles of dynein’s two motor domains are not tightly coordinated (DeWitt, Chang, Combs, & Yildiz,

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2012; Qiu et al., 2012). This raised intriguing questions regarding the motor’s ability to produce sustained forces that have since been addressed with single-molecule optical-trapping experiments.

1.2 Using an Optical Trap for Single-Motor Measurements A typical optical-trapping experiment is performed by bringing a bead that has been sparsely decorated with molecular motors in contact with the appropriate track (actin filament or microtubule). The track is usually firmly attached to the surface of the coverslip for the sake of experimental simplicity, though it may also be suspended in solution (Spudich, Rice, Rock, Purcell, & Warrick, 2011) if needed. As long as the bead is nearly spherical, the trapping laser applies no appreciable torque to it and the bead rotates freely until one of the motors on its surface engages with the track. It is critically important for the motors to be sparse on the surface of the bead since the experimenter has no other means for excluding cases with multiple motors simultaneously binding to the track. Once a motor engages with the actin filament or microtubule, it begins walking along it, pulling the bead away from the center of the trap. At small (under
100 nm) bead-trap separations, the optical trap acts as a Hookean spring, meaning that the restoring force pulling back on the motor is directly proportional to the distance by which it has pulled the bead out of the trap’s center. The bead-trap distance, and hence the force acting on the motor, is rapidly and accurately detected using back-focal plane interferometry (Gittes & Schmidt, 1998a), allowing for careful dissection of the motor’s mechanochemistry.

1.3 Measuring Motor Stall Forces The simplest trapping experiment, conceptually, is the measurement of a processive molecular motor’s stall force. The trapping laser is held in a fixed position such that the trapped bead is suspended just above the surface of the track. When a motor engages with the track and starts walking, the trap’s restoring force increases in parallel with bead-trap separation. Provided the trap is sufficiently stiff, the motor is unable to produce enough force to pull the bead completely out of the trap, and instead reaches an equilibrium position when the restoring force of the trap approximately equals the maximal force that the motor can produce in a forward step (Fig. 1). The force at which the net velocity of the motor becomes equal to zero is referred to as the stall force, and serves as a lower bound for the force produced in a single step of a motor. Since mechanical work is the product of force and displacement, knowledge of a motor’s step size, stall force, and number of

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Fig. 1 Stall force measurements of dynein and kinesin motors using a fixed trap. (A) Human kinesin-1 is attached to the anti-GFP antibody-coated bead via a C-terminal GFP fusion domain. (B) Kinesin walks toward the microtubule plus end, taking 8-nm steps until it stalls at
6 pN. The stall is marked with a red arrowhead. (C) Human dynein is complexed with the activating proteins dynactin and BICD2N and attached to the anti-GFP antibody-coated bead via an N-terminal GFP fusion domain. (D) The dynein–dynactin–BICD2N complex stalls at high forces comparable to those produced by kinesin-1. Note that typical dynein stalls are longer in duration than kinesin stalls. Figure adapted from Belyy, V., Schlager, M. A., Foster, H., Reimer, A. E., Carter, A. P., & Yildiz, A. (2016). The mammalian dynein–dynactin complex is a strong opponent to kinesin in a tugof-war competition. Nature Cell Biology, 18, 1019–1024. http://doi.org/10.1038/ncb3393.

ATP molecules hydrolyzed per step allows one to estimate the motor’s maximum energy efficiency. Kinesin and dynein are capable of producing
6 and
4 pN (Belyy et al., 2014, 2016; Svoboda & Block, 1994), respectively, while taking 8-nm forward steps. This means that despite vast structural differences, both motors approach
50% thermodynamic efficiency (free energy of single ATP hydrolysis is taken to be
80 pN nm). It is important that the stall force assay as described is only valid for highly processive motors whose typical runs in unloaded conditions (e.g., in a fluorescence motility assay) are at least several fold longer than the width of the potential well of the optical trap. Otherwise, the motor will frequently dissociate from the track before pulling the bead out to its maximal force, resulting in underestimation of the stall force.

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1.4 Force–Velocity Measurements Using Force Feedback The force–velocity (F–V) relationship of a motor yields detailed information about how external load affects its motility. Two motors with identical stall forces may exhibit dramatically different F–V curves depending on which step in the motor’s mechanochemical cycle is more sensitive to external load. For instance, myosin V readily walks backward under hindering loads, but does not accelerate past its unloaded forward velocity when it is pulled forward (Gebhardt, Clemen, Jaud, & Rief, 2006), resulting in a concavedown F–V curve. Meanwhile, yeast cytoplasmic dynein behaves in precisely the opposite way, effectively resisting hindering loads but rapidly walking forward when helped by an assisting force (Belyy et al., 2014). This behavior is immediately seen in dynein’s concave-up F–V curve (Fig. 2). Such different responses to load may arise from the tight coupling of specific steps of the motor’s enzymatic cycle to linear motion along the track. For instance, if a hypothetical motor is only capable of taking a step when an ATP molecule binds to one of its heads, while hindering external force decreases the likelihood of ATP binding (e.g., by closing the binding pocket), the motor’s velocity will be limited by ATP concentration under assisting loads and

Fig. 2 Force–velocity measurements of yeast cytoplasmic dynein. The solid black line   represents a fit to V ðF Þ ¼ V + eaFL=kT 1  eð△G + FLÞ=kT , where F is the external force, V+ is the unloaded velocity due to forward stepping only, ΔG is the energetic bias provided by ATP hydrolysis, L is the characteristic distance, and a is the dimensionless parameter that defines the partitioning of load-dependence between the forward and backward stepping rates. The net unloaded velocity V0 can be expressed as  

V0 ¼ V + 1  e△G=kT . Figure adapted from Belyy, V., Hendel, N. L., Chien, A., & Yildiz, A. (2014). Cytoplasmic dynein transports cargos via load-sharing between the heads. Nature Communications, 5, 5544. http://doi.org/10.1038/ncomms6544.

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approach zero under hindering loads. In addition to providing a window into the motor’s mechanochemistry, its F–V curve serves an important predictor of how the motor will respond to the balance of forces that may be encountered in the cell in the commonly occurring scenario of cargo transport by multiple motors. Typically, force–velocity measurements are performed by programming the trapping laser to follow a bead carried by a single motor, maintaining a constant bead-trap separation and therefore subjecting the motor to a constant load.

1.5 Studying Interhead Coordination Using Force-Dependent Unbinding Data Processive cytoskeletal motors comprise at least two distinct track-binding domains, and the F–V relationship lacks information about intramolecular forces that help the motor’s heads unite their pulling efforts. Understanding the effects of these intramolecular forces was of special interest in the study of dynein, because it does not strictly coordinate the stepping cycles of its two heads (DeWitt et al., 2012; Qiu et al., 2012). To measure rapidly and accurately the effects of intramolecular strain on a single head of a molecular motor, our lab developed a force-dependent unbinding assay and used it to demonstrate the inherent asymmetry in a single dynein head (Cleary et al., 2014). Briefly, a trapped bead decorated with monomeric motor heads is oscillated over the surface of a microtubule in a square-wave pattern. Occasionally, the head binds to the microtubule, and the trap is then held at a constant position until the head releases again. During this entire time period, the head feels a constant force from the trap. The experiment is almost fully automated and allows for the collection of thousands of unbinding events in a single assay. The unbinding times are then binned by force, and histograms are fitted to a single exponential decay to determine the microtubule unbinding rate of the motor in a given force range. The response of the motor’s head to a wide range of forces (Fig. 3) demonstrated that a dynein head releases from the microtubule in either an ATPdependent or an ATP-independent manner, explaining how the leading head of a full dynein motor may pull its trailing partner forward despite the apparent absence of interhead coordination. Moreover, we showed that a dynein head’s ATP-dependent release from the microtubule is inhibited by tension on the linker but is insensitive to force applied to the C-terminal domain, revealing a direct link between mechanical force on the head and its enzymatic cycle (Cleary et al., 2014).

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Fig. 3 Square wave unbinding force assay with yeast cytoplasmic dynein. (a) Polystyrene beads are sparsely decorated with dynein monomers. The trap oscillates between the two fixed positions A and B, spaced 250 nm apart along the microtubule’s axis. When a motor binds while the trap resides at position A (1), it resists the pull of the trap when the trap jumps to position B (2). During this time, the trap applies a constant force to the motor. After a period of time, the motor releases and the bead returns to the center of the trap (3). A real-time latching algorithm prevents the trap from returning to position A while the motor remains bound. (b) The interplay between dynein’s ATPase cycle and intramolecular forces can be studied by attaching the optically trapped bead either to the mechanically active linker domain (top) or the rigid C-terminus (bottom) of the monomer. (c) Release rates of dynein monomers depend on both force and nucleotide concentration. When pulled from the C-terminus, dynein releases more rapidly in the presence of ATP than in the ATP-free (apo) case. However, applying the force from the linker eliminates the ATP-dependent increase in release rates, demonstrating that force felt through the linker regulates dynein’s ATPase cycle. In all cases, dynein releases more rapidly when pulled toward the microtubule minus end. Figure adapted from Cleary, F. B., Dewitt, M. A., Bilyard, T., Htet, Z. M., Belyy, V., Chan, D. D., … Yildiz, A. (2014). Tension on the linker gates the ATP-dependent release of dynein from microtubules. Nature Communications, 5, 4587. http://doi.org/10.1038/ncomms5587.

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2. EXPERIMENTAL SETUP AND TROUBLESHOOTING 2.1 Optical Layout of the Trap Our trapping microscope consists of a 2-W 1064-nm fiber-coupled CW laser (Coherent Compass) for trapping beads, a pair of acousto-optical deflectors (AODs; AA Opto Electronic DTSXY-400) for precise beam steering, and a 1.49 NA oil immersion objective (Nikon CFI Apo TIRF 100 ) which focuses the trapping beam to a nearly diffraction-limited spot in the image plane. The trap utilizes back-focal plane interferometry for position detection (Gittes & Schmidt, 1998a; Toli
c-Nørrelykke et al., 2006) by imaging the trapping laser’s beam onto a 200-kHz position sensitive detector (PSD; First Sensor DL100-7-PCBA3). Fluorescence laser lines are available for TIRF imaging, and a separate 845-nm solid-state laser is reflected off the coverslip and projected onto a separate PSD for long-term focus stabilization. Precise power control is achieved by discarding the required percentage of the laser’s output into a beam dump, as determined by a half-wave plate mounted on a motorized rotary mount followed by a polarized beam-splitter cube. Several excellent reviews and guides have been written on the general layout and calibration of optical traps (Bustamante, Chemla, & Moffitt, 2009; Neuman & Block, 2004). We will instead focus on often-overlooked aspects of optical trap design—its custom-written control software and sources of experimental noise in trapping measurements.

2.2 Software Control of the Trap Good software development practices are key to building a robust and reliable scientific instrument such as an optical trap. A minimal effort, “just get it done” approach quickly results in ballooning development and troubleshooting times as the project increases in complexity. The standard tool for scientific hardware control is LabVIEW, a graphical data flow language that comes packaged with a comprehensive library of instrument drivers and is relatively easy to start coding in for someone with no prior programming experience. However, the design of the language makes it easier for an inexperienced programmer to turn the simplest program into an unreadable tangled mess of wires, giving new meaning to the expression “spaghetti code.” A large instrument control suite written in this style not only becomes

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impossible to debug, but also carries a risk of undetected persistent errors making their way into the acquired data files and compromising the validity of the experimental results. There are several important practices which will ensure that the code remains readable and amenable to debugging as it grows. First, all code should be subdivided into logical subfunctions (termed subVIs in LabVIEW), each of which should ideally fit on a single monitor screen. Each subVI should carry out one specific task on a small number of inputs. If the programmer cannot describe in one or two sentences exactly what a given subVI does, its code is probably not logically partitioned. Proper subdivision also helps greatly with debugging: if a subVI’s expected behavior can be succinctly explained, it becomes easy to test whether it performs as expected by feeding a sample input into it and checking whether the output exactly matches the programmer’s expectation. Second, because LabVIEW’s data structures (wires) represent data flow rather than data storage, care must be taken from the start to organize and bundle data flow. Perhaps the most useful, and underutilized, organizational feature of LabVIEW is wire clustering. If several pieces of information are used to describe the state of some logical entity (e.g., a photosensitive detector might simultaneously output four channels of data), there is no reason to have four separate wires run across the screen from one subVI that processes detector data to the next. Bundling-related data makes the program easier to follow, troubleshoot, test, and update. Finally, LabVIEW’s clunky code annotation system and lack of variable names per se puts an extra burden on the programmer to document the program in a way that is easy for others to follow. The best way to ensure readability is to subdivide the program into manageable logical chunks and connect the chunks using a few well-placed clustered wires. The last step toward readability is to assign meaningful names to all input and output terminals of the individual subVIs. Note that every LabVIEW wire inherits the name of its source terminal, meaning that properly labeling the outputs of subVIs will automatically result in naming all wires running across the block diagrams. The end result is a top-level block diagram that fits on a single monitor screen and whose general logic flow can be parsed by following the names of subVIs and the relatively few high-level cluster wires connecting them.

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2.3 Noise Elimination Precise measurement of nanometer-scale distances is often an absolute requirement for step size measurements of molecular motors using optical traps. Therefore, identifying the smallest sources of noise and eliminating ones not inherent to the experiment becomes a key task for any researcher designing or troubleshooting the instrument. Before embarking on the noise hunt, it is critical to understand the properties and fundamental limitations of thermal (or Brownian) noise, which sets fixed bounds on the spatial and temporal resolution achievable in a trapping experiment (Bobroff, 1986; Gittes & Schmidt, 1998b). Other, non-Brownian, sources of noise can generally be identified and either removed, minimized, or corrected for. Generally, the easiest sources of noise to identify are those with a welldefined characteristic frequency. By immobilizing beads on the surface of a coverslip, focusing the trap on one of the beads, recording a position trace over several seconds, and taking the trace’s Fourier transform, a compound power spectrum of all the contributing non-Brownian noise sources could be obtained. Single-frequency noise will appear as a sharp, distinct spike in this power spectrum. One real example of such noise in our trap came from a cooling fan built into a camera connected to the microscope body. When the fan is turned on, we observed a clear spike at the fan’s rotational frequency; the solution was to disable the fan and water cooling the camera. Another common source of single-frequency noise is “mains hum” or leakage of the 60 Hz AC current signal from the building’s power supply into the trap’s sensitive detection electronics. This leakage can either occur directly, e.g., as a result of using a low-quality AC-to-DC converter to power the detector, or through an unexpected path (for instance, if ceiling lights are on in the room during recording, their light output will also oscillate at 60 Hz and trigger periodic currents in the detector). The two examples earlier can be solved by using higher-quality AC-to-DC converters and performing the experiments with room lights turned off, respectively. Unfortunately, many sources of noise and drift exhibit complex power spectra and cannot be so easily traced to a specific source. Examples include, but are certainly not limited to, stage drift, laser pointing instabilities, vibrations in the microscope body, electrical leaks and faults in the detection electronics wiring, ground loops, temperature drift due to differences in thermal expansion coefficients of individual microscope parts, unstable mounting of the sample chamber, and small liquid currents inside the sample chamber. Troubleshooting an elusive noise source can be a frustrating experience,

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and in many cases, a better solution to eliminate the contribution of all of these noise sources is to measure the position of a fiducial marker separately and subtract it from the original trace. In dual-beam trapping experiments, which are already decoupled from the surface, most noise is canceled out and the remaining noise originates from the short differential segments where the beams making the two traps do not travel along the same path (Bustamante et al., 2009). The differential path can be eliminated entirely by creating both traps, individually steerable along both x and y directions, with the same laser using a pair of AOD’s. Commanding the AOD’s to move rapidly the laser beam between the target positions, one obtains multiple time-shared traps (Visscher, Gross, & Block, 1996) that use the same optical path and the same detector. When the position of the first trapped bead is subtracted from the position of the second, any external noise sources present in both signals nearly disappear. The concept is fully applicable to surface-coupled experiments, though it necessitates an additional step of immobilizing fiducial markers on the surface of the coverslip (Carter et al., 2007). Should higher-resolution surface-coupled experiments be required in surface-coupled trapping measurements, we highly recommend implementing fiducial-based noise correction, for example, as reviewed in Perkins (2014), instead of hunting for ever smaller sources of noise in the instrument. In principle, subnanometer resolution can be achieved in a noise-corrected optical trap over the timescale of minutes at sampling frequencies of several hundred Hertz (Perkins, 2014).

3. EXPERIMENTAL PROTOCOLS 3.1 Coating Polystyrene Beads With Antibodies 3.1.1 Stock Reagents and Solutions 1. Carboxylated latex microspheres/beads,
1 μm in diameter (e.g., 4%, w/v from Life Technologies) 2. Activation buffer (10 mM MES, 100 mM NaCl, pH 6.0) 3. 1 PBS (phosphate-buffered saline), pH 7.4 4. 10% Sodium azide in ddH2O 5. EDC ((1-ethyl-3-(3-dimethylaminopropyl)carbodiimide hydrochloride) crosslinker (hygroscopic powder, stored in desiccator at 20°C) 6. NHS-S (N-hydroxysulfosuccinimide) crosslinker (hygroscopic powder, stored in desiccator at 20°C) 7. DMF (Dimethylformamide, hygroscopic)

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8. Antibodies of your choice (e.g., 500 μL of 0.5 mg/mL, adjust other volumes accordingly) 9. BSA (bovine serum albumen) 50 mg/mL in ddH2O 3.1.2 Protocol 1. Add 200 μL of beads to 800 μL of activation buffer. 2. Spin at 6000 rcf for 3 min, discard supernatant. 3. Resuspend pellet in 800 μL of activation buffer by pipetting and vortexing. 4. Repeat steps 2 and 3 twice to wash the beads. 5. Dissolve 1 mg of EDC in 1 mL of DMF and 1 mg NHS-S in 2 mL of DMF. 6. Add 20 μL of the dissolved EDC and 40 μL of dissolved NHS-S to the beads. 7. Sonicate for 2–3 min or until visible clumps disappear, then nutate or vortex at low speed for 30 min. 8. Rinse three times in PBS by spinning, resuspending, and vortexing. 9. Check the beads under 100 magnification in a bright-field microscope. If they are clumpy, sonicate for 2–5 min and check again. 10. Spin the beads down and resuspend the pellet in 800 μL of PBS. 11. Add the
200 μL of 0.4 mg/mL antibody to the 200 μL of beads. 12. Nutate or shake for 30 min. 13. Add 100 μL of 50 mg/mL BSA in ddH2O and nutate for 2–4 h at room temperature (or O/N at 4°C) to passivate the beads. 14. Rinse three times in PBS, resuspend by pipetting and vortexing—do NOT sonicate the beads for longer than 10 s once they are mixed with the antibody. 15. Resuspend in four times the original volume of beads with 0.1% azide and 0.5 mg/mL BSA (i.e., 784 μL PBS + 8 μL 50 mg/mL BSA + 8 μL 10% azide). 16. Store at 4°C. Never freeze the beads. 3.1.3 Notes Spin times and speeds will vary depending on the diameter of beads used. For 0.9–1.0-μm diameter beads, each spin should be
3 min at 6000 rcf (
8000 rpm in a tabletop centrifuge); for 0.2 μm beads, 5 min at 16,000 rcf (
13,000 rpm). Adjust the spins as required to ensure that most beads go down into the pellet, but the pellet is not too hard to avoid their aggregation.

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Addition of excess EDC and NHS-S leads to irreversible clumping of the beads. The amount of crosslinker to be added to the beads should be adjusted depending on the total surface of the beads in the mixture. More information can be found from the instructions given by the beads’ manufacturer.

3.2 Human Dynein Optical Trapping Sample Preparation 3.2.1 Stock Reagents and Solutions 1. ATP (100 mM or as desired) 2. Human cytoplasmic dynein (
1 μM) tagged with GFP at the N-terminus 3. Dynactin (
1 μM) 4. BICD2N (
1 μM) 5. Oxygen-scavenging system: a. Protocatechuate dioxygenase (PCD, 3.5 mg/mL(100)) b. Protocatechuic acid (PCA, 250 mM (100)) 6. 1,4-Dithiothreitol (DTT, 1 M) 7. Sea urchin axonemes (Gibbons & Fronk, 1979) labeled with Cy5, stored at 20°C in 40% glycerol 8. Casein (40 mg/mL) 9. Motility buffer (MB) (30 mM HEPES, 5 mM MgSO4, 1 mM EGTA, pH 7.0 with KOH) 10. α-GFP antibody-coated polystyrene beads 3.2.2 Solutions to Make Fresh for Each Sample MBC (Motility buffer with casein) 1. Casein (40 mg/mL stock): 10 μL 2. MB: 140 μL AXO (Axoneme solution) 1. Axoneme stock: 1 μL 2. MB: 9 μL Stepping buffer (adjust [ATP] as needed. This recipe makes 2 mM ATP) 1. Casein (40 mg/mL stock): 3 μL 2. PCA stock: 1 μL 3. PCD stock: 1 μL 4. ATP stock: 2 μL 5. DTT stock: 1 μL 6. MB: 2 μL

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3.2.3 Protocol 1. Dilute the dynein stock in MBC to make
0.2 μM dynein. 2. Dilute the dynactin stock in MBC to make
1 μM dynactin. 3. Dilute the BicD2N stock to make
0.4 μM BicD2N. 4. Mix dynein with dynactin and BICD2N at a 1:5:2 molar ratio and incubate on ice for 5 min (1 μL diluted dynein + 1 μL diluted dynactin + 1 μL diluted BICD2N). 5. Prepare the sample chamber by affixing two parallel pieces of doublesided tape (Scotch 3136) onto a glass microscope slide, leaving a
3 mm gap in the middle, then pressing a #1 coverslip onto them and peeling off any remaining tape. 6. Prepare the motor dilutions in MBC. 7. Mix bead stock by pipetting, take out 1.5 μL into a small tube, sonicate for 7 s. 8. Mix the 1.5 μL sonicated beads with 2 μL of diluted protein; let it sit for 10 min on ice. 9. After 10 min, add 14 μL of MB and 2 μL of stepping buffer to the motors + beads solution. 10. Flow AXO into the sample chamber and wait for 30 s. 11. Wash slide with 40 μL MBC and let it sit for at least 3 min. 12. Flow the 20 μL of Motors + Beads + MB + Stepping buffer solution prepared in steps 7 and 8. 13. Seal ends of sample chamber with nail polish.

3.3 Trap Position vs AOD Command Signal Calibration 3.3.1 Protocol 1. With the trapping laser off, mount a reticle onto the XY stage and determine the conversion factor between camera pixels and distance in the image plane (a typical convenient range is between 50 and 100 μm per pixel). 2. Remove the reticle and mount a flow cell containing a very dilute bead solution. 3. Turn on the trapping laser and trap a single bead. 4. Divide the allowed frequency range of the AOD into 5 to 10 equally spaced increments (e.g., for an AOD with a 75 MHz central frequency and a 60–90 MHz range, you may choose to test multiples of 5 MHz: 60, 65, 70, 75, 80, 85, and 90 MHz). 5. Set the y-axis of the AOD to the central frequency.

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6. Send the chosen frequencies to the x-axis AOD one by one, causing the bead to move across the AOD’s allowed range. Save the camera image at each position. 7. Repeat the scan for the y-axis of the AOD, keeping the x-axis at the central frequency. 8. For each resulting image, determine the x and y coordinates (in pixels) for the center of the trapped bead at each commanded AOD frequency. 9. For each axis, plot the bead position vs commanded frequency. The points should all neatly fall on a line. The slope of this line is the desired AOD calibration coefficient that allows you to convert between the AOD command signal, in megahertz, and the trap position in the image plane. Please note that if the axes of your AOD are not perfectly aligned with the x- and y-axes of the camera, you may need to rotate the signal accordingly for the calibration to remain valid. 3.3.2 Note This calibration only needs to be performed after major microscope realignment. There is no need to repeat it on a daily basis as both camera pixel size and AOD frequency vs displacement remain very nearly constant.

3.4 Trap Stiffness and PSD Response Calibration 3.4.1 Protocol 1. Trap a single bead, bring it to center. 2. Find the surface of the coverslip and position the bead just above it (not above an axoneme). 3. Move the bead up 3 μm into solution. 4. Record bead position for 20 s with at least 20 kHz sampling frequency. Make sure that a single bead remains trapped. If a second bead comes into the trap, discard it and repeat with a new single bead. 5. Decrease the laser power from your working level to about 5 mW. This decreases the trap’s corner frequency, which is critically important for raster scan (see note later). 6. Perform an AOD raster scan along the x-axis to determine the PSD response coefficients (i.e., the conversion factors between dimensionless PSD units and nanometers; see note later). 7. Repeat the AOD raster scan along the y-axis. 8. Slowly move the focus down 3 μm to bring the bead back to the surface. 9. Repeat the AOD raster scan for both x and y axes.

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10. Use the PSD response coefficients collected at the surface for interpreting trap data. 11. Determine trap stiffness via the power spectrum method (Toli
cNørrelykke et al., 2006) using the 20-s file collected in step 4 and the PSD response coefficients found in step 6. 3.4.2 Note on Raster Scanning A key feature of back-focal plane interferometry is that the PSD signal is sensitive only to the distance between the center of the bead and the center of the trap. The trap’s true position is known at any point in time thanks to the AOD command signal calibration described in the previous section. The raster scan in steps 6, 7, and 9 above thereby amounts to repeatedly commanding the trap to jump away from the central position for a brief period of time, measuring the corresponding PSD response, and commanding the trap to return to the central position before the bead reacts and moves away from the center by an appreciable distance. To avoid such unwanted bead movement, the trap must remain in the central position for the majority of time (in our case, 98% of the time) and the sampling frequency must be at least an order of magnitude higher than the trap’s corner frequency. For instance, in a typical raster scan in our trap, the corner frequency is
100 Hz, while the sampling frequency is 20 kHz. The trap remains in the central position for 98 data points (4.9 ms), and then moves to the test position for 2 data points (0.1 ms). The trap’s low corner frequency ensures that the bead does not have time to react to the change in trap position and that we measure the true PSD response for each tested trap-bead separation. We typically test 20 distinct trap-bead separations ranging from 300 to +300 nm and average the outcomes of 20 raster scans to eliminate most thermal noise and construct a clean PSD calibration curve. Also note that we perform two sets of raster scans, one 3 μm away from the surface and one directly at the surface. The reason behind this is that when using a high-NA oil immersion objective and an oil immersion condenser, the PSD response changes significantly as a function of trap distance away from the surface. Meanwhile, trap stiffness remains nearly independent of this distance, provided that the objective is operating within the limits of its working distance. We therefore use the coefficients from the scan performed 3 μm from the surface to convert raw PSD data to distances in the stiffness calibration recording (which must be performed at least two bead diameters from the surface to avoid surface effects). Meanwhile, the coefficients obtained from the surface-level scan are used to convert experimental data to distances.

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3.5 Dynein Stall Force Measurements 1. 2. 3. 4. 5.

6. 7. 8.

9. 10.

11.

Load prepared sample onto the trap microscope. Find the surface of the coverslip and focus 2–3 μm above it. Catch a single bead with the trapping laser. Calibrate the trap as described in the previous section. Identify a fluorescently labeled axoneme (if bright-field illumination is very well aligned, axonemes may be visible in the absence of fluorescent markers). Steer the trapping laser back to its central position. Move the stage such that the bead is now trapped directly above the axoneme. By moving the objective down (i.e., changing the microscope’s focus), very slowly lower the bead onto the axoneme until the moment when it just barely touches the axoneme’s top surface. It is easiest to determine when this happens by looking at the total intensity of the trapping laser’s light impinging on the PSD. This intensity stays very nearly constant when the bead is trapped in solution, but as the bead touches the surface the intensity changes substantially because the bead moves slightly out of the trap’s focus and refracts a different percentage of light passing through it. With the bead positioned just above the surface of the axoneme, wait
30 s to see if the bead binds to the axoneme or begins walking. If the bead exhibits any activity, begin recording a full data file. If it does not, release it by turning off the trap (this also verifies that the bead is not stuck to the surface) and repeat steps 4 through 8 with a different bead. Make sure that 30% of them are sticking to axonemes, your beads may be poorly passivated during labeling or you may be suffering from poor motor protein quality. We recommend checking the motility using a single-molecule fluorescence assay before troubleshooting the trapping assays.

3.6 Identification of Motor Stalls In a fixed trap assay, every type of motor exhibits different stall behavior. For example, dynein walks out to approximately the maximal force of
4 pN and then remains tenaciously bound to microtubules for as long as a minute

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or two, taking occasional forward and backward steps, until it finally releases from the track and the bead snaps back into the center of the trap. On the other hand, kinesin-1 does not stall for a long time. Instead, when the motor reaches its maximal force, it remains bound to the microtubule for only a bit longer than the average duration of the step and releases from the microtubule in response to the resistive load from the trap. Before collecting stall statistics, it is essential to perform preliminary experiments over a range of trap stiffness values and find a stiffness at which motor-driven bead displacements cover a significant percentage of the trap’s linear range (
75 to 100 nm), but no beads pull strongly enough to escape from the trap altogether. This is an important compromise, as increasing the trap stiffness greatly reduces the experimental noise, but using a trap that is too stiff for the application will result in motors taking only a few steps before stalling and will increase the uncertainty in stall force assignment. On the other hand, an overly weak trap will allow the strongest motors to escape, causing the measured stall force to be underestimated. It is also essential to define a working criterion for what constitutes a stall. Typically, this must be a pause in motility that lasts several times longer than the motor’s mean step duration, followed by a clean fallback of the bead into the trap center. For instance, the metric we typically use for dynein is that a stall must last longer than 0.5 s, followed by a clean return to center, meaning that the bead does not first fall back to an intermediate position (this could mean that a second motor is present or that the bead is not properly passivated). Please note that a weakly processive motor may not allow one to determine the stall force accurately. If the motor repeatedly releases from the microtubule without reaching a plateau, its stall force cannot be accurately measured using the technique described here.

3.7 Force-Feedback Assay 1. Load the sample and position a bead just above an axoneme as described earlier. 2. If the bead exhibits motility, arm the force feedback routine and begin recording data. 3. Our feedback algorithm is written such that once it is armed, it will only engage once the bead is pulled out of the center of the trap by more than a user-defined threshold value. This is done because performing feedback on a bead that is not firmly attached to the axoneme via a molecular

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motor causes large-scale position fluctuations. The position threshold serves to ensure that a motor is engaged with the track and actively pulling the bead before feedback is attempted. Furthermore, motors have a strictly preferred polarity (e.g., dynein walks toward the minus ends of microtubules), and the feedback algorithm assumes that the direction in which the threshold was reached is the preferred direction of the motor. 4. You may collect several runs from the same bead; however, be wary of skewing the data toward a small subset of not necessarily representative highly active beads. After you have collected a few runs, release the bead even if it continues to walk and move on. 5. Calibrate the trap at the end of every sample’s lifetime.

3.8 Construction of Force–Velocity Curve To build a force–velocity curve for a molecular motor, several runs in forcefeedback mode must be collected at each force being tested. When deciding which forces to test, choose evenly spaced values that extend at least 25–30% past the motor’s stall force in the hindering direction and half of the stall force in the assisting direction (e.g., for a motor that stalls at 3 pN, at the minimum test the interval between 4 pN hindering and 2 pN assisting) (Fig. 4). The experiments may be time consuming if the motor is not well behaved, so test a few forces evenly spaced throughout the entire range first and then decide whether a more detailed dataset is warranted. The definition of a valid run should be established for a given motor. A typical definition of a run in our assays is: 1. Force feedback must be engaged for at least half a second. 2. The bead must travel over 50 nm over the course of the run. 3. During the run, the bead may not take steps larger than the motor’s maximal measured step size, as such “steps” may indicate release and rebinding events. Once a sufficient number of runs is collected from independent beads, the mean velocity of each run is determined by either taking the slope of the line connecting the start and end points or fitting the entire run to a line. The mean velocity for each force is plotted in a F–V curve, which can then be fitted to a simple single-state motor model to determine whether the forward or backward stepping rate of the motor is more sensitive to external load (Belyy et al., 2014).

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Fig. 4 Representative force feedback traces from human dynein with the adapter proteins dynactin and BICD2N. (A) When the force is clamped at super-stall forces (6 pN), dynein walks backward in response to the constant pull of the trap. (B) At a low feedback force of 0.4 pN, dynein walks rapidly toward the minus end of the microtubule despite the trap’s hindering pull. Figure adapted from Belyy, V., Schlager, M. A., Foster, H., Reimer, A. E., Carter, A. P., & Yildiz, A. (2016). The mammalian dynein–dynactin complex is a strong opponent to kinesin in a tug-of-war competition. Nature Cell Biology, 18, 1019–1024. http://doi.org/10.1038/ncb3393.

3.9 Square Wave Unbinding Force Assay 1. Prepare polarity marked microtubules or axonemes since there is no other way to determine track polarity in the square wave experiment. A quicker and dirtier method (though still very effective) is to use unlabeled tracks and flow fluorescently labeled motors of known polarity (e.g., GFP-dynein) into the sample. The motors will accumulate at one tip of the track, the microtubule minus end in the case of dynein, and will serve as a fluorescent polarity marker. 2. Coat the beads with motor monomers instead of fully assembled active dimers as in the previous assays. 3. Load the prepared sample onto the trap microscope. 4. Catch a bead and bring it in contact with an axoneme. Define the angle of the axoneme in the software to tell the trap which axis the square wave should follow. 5. Engage the square wave routine. This will move the trap between two predefined positions several hundred nanometers apart (typically 300 nm). The period of the square wave can be any arbitrary number since it does not impact the outcome of the experiment; we typically use a 2 Hz wave.

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6. It is important for the square wave algorithm to detect a binding event and keep the trap stationary until the motor releases from the microtubule. This is easily done by monitoring the running mean displacement of the bead from the center of the trap and define a binding event as any >100 ms window wherein bead-trap separation remains greater than some reasonable value such as 10 nm. The exact settings will vary somewhat depending on the trap stiffness used in the experiment.

3.10 Analysis of Unbinding Force Data Data used to determine the unbinding force consist of a large number of periodic square waves with occasional interspersed binding event (Fig. 3). A binding event is apparent when the bead remains in an intermediate position instead of following the trap on the up- or down-swing of the square wave. We use a semiautomated algorithm for identifying such events and fitting them to a single-step function. Individual step fit results are quickly scanned by eye and ones with more than one stepwise transition, position drift, or excessively high noise levels are removed from further analysis (these can all indicate bead stickiness or the simultaneous binding of multiple motors). Resulting binding events are then binned by force and their durations plotted on a single graph. An important consideration is to ensure that the polarity of the track is noted and correctly marked in each data file, as in contrast to stall and force–velocity assays the information about which direction is “forward” for the motor cannot be easily determined from the data file itself. A unique strength of this approach is that a single assay simultaneously probes a large force range, with the limits of the range being determined by the trap’s spring constant and square wave amplitude.

4. CONCLUSION As we have discussed in this chapter, optical trapping remains a powerful tool for dissecting the mechanochemical properties of processive motor proteins. The information that can be obtained in trapping experiments ranges from basic motor parameters such as maximal force production and response to external load to detailed insight into the coupling between ATPase activity of a single motor head and the force applied to it by its partner head (Cleary et al., 2014). While it may appear that dynein, kinesin, and myosin have been thoroughly investigated, recent studies indicate that many adapter proteins regulate cytoskeletal motors by directly modifying their force production and processivity (Belyy et al., 2016; McKenney et al.,

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2014; Schlager et al., 2014). Furthermore, the mechanisms underlying cargo transport by large teams of molecular motors still remain controversial (Hancock, 2014; Hendricks et al., 2010; Leidel, Longoria, Gutierrez, & Shubeita, 2012; Soppina, Rai, Ramaiya, Barak, & Mallik, 2009). Optical trapping will remain an ideal tool for studying the mechanochemical effects of motor protein regulation and motor teamwork in the years to come.

ACKNOWLEDGMENTS This work was supported by the NIH (GM094522 (A.Y.)), an NSF CAREER Award (MCB-1055017 (A.Y.)), and an NSF Graduate Research Fellowship (DGE 1106400 (V.B.)).

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CHAPTER THREE

Single-Molecule Optical-Trapping Techniques to Study Molecular Mechanisms of a Replisome B. Sun*, M.D. Wang†,{,1 *School of Life Science and Technology, ShanghaiTech University, Shanghai, PR China † Laboratory of Atomic and Solid State Physics, Cornell University, Ithaca, NY, United States { Howard Hughes Medical Institute, Cornell University, Ithaca, NY, United States 1 Corresponding author: e-mail address: [email protected]

Contents 1. Introduction 2. Instrument Design, Experimental Configuration, and Sample Preparation 2.1 Layout of an Optical-Trapping Apparatus 2.2 Experimental Configuration 2.3 Preparation of Experimental Sample Chambers 3. Molecular Mechanisms of Individual Proteins in the Replisome Revealed by Optical-Trapping Techniques 3.1 DNA Template Design and Construction 3.2 Helicase-Unwinding Assay 3.3 Helicase Translocation Assay 3.4 Polymerase Strand Displacement Assay 4. Single-Molecule Studies of the Response of a Replisome to DNA Damage 4.1 DNA Template Design and Construction 4.2 Response of Individual Proteins to DNA Damage: Helicase-Unwinding Assay and Polymerase Strand Displacement Assay. 4.3 Response of a Replisome to DNA Damage: Leading-Strand Replication Assay 5. Data Analysis 5.1 DNA Elastic Parameter Determination 5.2 DNA Extension Conversion to Number of Base Pairs 5.3 Helicase Slippage Determination and Processivity Measurements 5.4 Unwinding Rate and Replication Rate Measurements and Comparison 5.5 Fate of a Replisome Encountering DNA Damage 6. Unique Features of the Bacteriophage T7 Replisome Revealed by Single-Molecule Optical-Trapping Techniques 6.1 Helicase Slippage and Subunit Coordination 6.2 Inherent Tolerance of the T7 Replisome to DNA Damage

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Abstract The replisome is a multiprotein molecular machinery responsible for the replication of DNA. It is composed of several specialized proteins each with dedicated enzymatic activities, and in particular, helicase unwinds double-stranded DNA and DNA polymerase catalyzes the synthesis of DNA. Understanding how a replisome functions in the process of DNA replication requires methods to dissect the mechanisms of individual proteins and of multiproteins acting in concert. Single-molecule optical-trapping techniques have proved to be a powerful approach, offering the unique ability to observe and manipulate biomolecules at the single-molecule level and providing insights into the mechanisms of molecular motors and their interactions and coordination in a complex. Here, we describe a practical guide to applying these techniques to study the dynamics of individual proteins in the bacteriophage T7 replisome, as well as the coordination among them. We also summarize major findings from these studies, including nucleotide-specific helicase slippage and new lesion bypass pathway in T7 replication.

1. INTRODUCTION DNA replication is carried out by the replisome, a large protein complex that includes DNA helicase, DNA polymerase (DNAP), and other proteins (Benkovic, Valentine, & Salinas, 2001). The bacteriophage T7 replisome has been identified to be a simple and efficient model system for the study of DNA replication (Hamdan & Richardson, 2009), as only four proteins are required for basic phage replication, yet it mimics more complex systems. Briefly, T7 DNAP, a 1:1 complex of gene 5 protein (gp5) and its processivity factor Escherichia coli thioredoxin (trx), is responsible for nucleotide polymerization (Tabor, Huber, & Richardson, 1987). The protein product of T7 gene 4 (gp4) provides helicase and primase activities (Matson, Tabor, & Richardson, 1983). The helicase activity, required for unwinding the parental DNA strand, resides in the C-terminal half of the protein, while the primase activity, required to initiate lagging-strand DNA synthesis by synthesizing RNA primers, is located in the N-terminal of the protein. T7 helicase is a hexameric motor and couples the hydrolysis of nucleoside triphosphate to translocate along single-stranded DNA (ssDNA) and unwind double-stranded DNA (dsDNA) (Donmez & Patel, 2008; Singleton, Dillingham, & Wigley, 2007). Finally, the product of gene 2.5 of bacteriophage T7 (gp2.5), a single-stranded DNA-binding (SSB) protein,

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coats ssDNA to remove its secondary structure (Kim & Richardson, 1993). Although tremendous advances are being made in our understanding of the structures and functions of these proteins (Hamdan & Richardson, 2009), their dynamic and mechanistic properties are not fully understood. For example, how do different subunits of the hexameric helicase coordinate their chemical and mechanical activities to translocate along ssDNA? How does the helicase coordinate with DNAP during replication? How does DNAP deal with DNA lesions in a template strand? These questions are well suited to be investigated by optical trapping, a powerful single-molecule technique that offers the unique ability to observe and manipulate biomolecules at the single-molecule level. By directly revealing biomolecular dynamic behaviors in real time, this approach can provide essential information and complement ensemble studies in understanding various biological systems (Moffitt, Chemla, Smith, & Bustamante, 2008; Sun & Wang, 2016). Optical trapping provides flexible control of both force and extension of the substrate, enabling rapid switching between different modes of operation. In this chapter, using the bacteriophage T7 replisome as an example, we detail experimental procedures utilizing optical-trapping techniques to study molecular mechanisms of individual proteins in the replisome. In addition, we also provide a practical guide to dissecting the response of the T7 replisome to DNA damage.

2. INSTRUMENT DESIGN, EXPERIMENTAL CONFIGURATION, AND SAMPLE PREPARATION 2.1 Layout of an Optical-Trapping Apparatus In general, an optical trap is generated by using a high-numerical aperture microscope objective to focus tightly a laser beam to a diffraction limited beam waist. The gradient of the intensity provides the trapping force, which can be used to manipulate a trapped dielectric microsphere. Here, we use a single-beam optical-trapping instrument containing the minimal set of optical components required for the operation of a high-precision instrument of its kind (Fig. 1) (Brower-Toland & Wang, 2004; Koch, Shundrovsky, Jantzen, & Wang, 2002; Li & Wang, 2012). In brief, a 1064-nm Gaussian laser ( J20I-8S-12K-NSI, Spectra-Physics Lasers, Inc., Mountain View, CA) is coupled to a single-mode fiber (PMJ-A3A, 3AF-1064-6/125sAS-12-1, Oz Optics, Carp, ON). After collimation, the beam is expanded by a telescope lens pair and then focused onto the back focal plane of a 100
, 1.4 NA oil immersion microscope objective (Plan Apo 100
/1.40 Oil IR, Nikon, Melville, NY). An acousto-optic deflector (AOD) (AOBD

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Fig. 1 Layout of the optical-trapping apparatus and a schematic of the sample chamber. See text for a detailed description of the setup.

N45035-3-6.5DEG-1.06, NEOS Technologies, Inc., Melbourne, FL) is placed between the laser aperture and the beam expander to adjust the laser intensity. The beam is then introduced into a modified Eclipse Nikon TE200 inverted microscope’s imaging path (Nikon, Melville, NY). Upon beam collection by a condenser, the laser beam is imaged onto a quadrant photodiode (S5981, Hamamatsu, Bridgewater, NJ), at which a deflection voltage signal reflects a displacement of a trapped microsphere. A highprecision 3D piezoelectric stage (Nano-PDQ350HS, Mad City Labs, Madison, WI) is used to position a sample chamber. Analog voltage signals generated by the position detector and piezo stage position sensor are antialias filtered at 5 kHz (part number 3384, Krohn-Hite, Avon, MA) and digitized at 7–13 kHz, using a multiplexed analog-to-digital conversion PCIe board (NI PCIe-6259, National Instruments Corporation, Austin, TX). The instrument

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calibration methods and experimental control modes, such as force clamp and velocity clamp, were described in previous publications (Li & Wang, 2012).

2.2 Experimental Configuration To mimic a DNA replication fork, we typically start an experiment with a DNA template containing a fork junction initially or after mechanically unzipping of a dsDNA (see Sections 3.1 and 4.1) ( Johnson, Bai, Smith, Patel, & Wang, 2007; Sun et al., 2011, 2015). As shown in Fig. 2, the ends of the leading and lagging strands are differentially labeled, generally by digoxigenin and biotin. One strand is attached to a trapped 500 nm microsphere via a biotin/streptavidin connection, and the other strand is anchored to a microscope coverslip surface via a digoxigenin/antidigoxigenin connection. The microsphere is held in a feedback-enhanced optical trap so that both its position and force can be measured. Experiments are typically conducted under a constant force where the coverslip position is modulated to maintain a constant force on the trapped microsphere. Helicase unwinding, DNA synthesis or degradation, and helicase-unwinding coupled DNA synthesis are reflected as a change in the DNA length between the microsphere and the anchor point on the coverslip.

Fig. 2 Experimental configuration for the studies of a replisome. An optical trap is used to exert a force on a trapped microsphere and monitor extension change resulting from the activities of motor proteins at the fork. The extremities of a DNA template are attached to the coverslip and microsphere via digoxigenin/antidigoxigenin and biotin/streptavidin connections, respectively.

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2.3 Preparation of Experimental Sample Chambers Prior to mounting onto an optical setup, sample chambers are prepared at room temperature in a humid chamber to minimize buffer evaporation ( Johnson et al., 2007; Li & Wang, 2012). Briefly, two thin pieces of double-stick transparent tape (0.1 mm thick) are first applied in parallel with 5 mm separation to a coverslip (24 mm
40 mm
0.15 mm). Then, a glass slide (25 mm
76 mm
1.2 mm) is perpendicularly placed on top of the coverslip, creating a 15 μL volume channel bordered by the two strips of tape (Fig. 1). To immobilize individual DNA tethers in the sample chamber, different solutions are sequentially flowed into the chamber. Polyclonal sheep antidigoxigenin (Roche Applied Sciences), diluted in PBS buffer, is used first to coat the coverslip, followed by a blocking buffer (a typical blocking agent is casein sodium salt from bovine milk, Sigma-Aldrich Co.) to prevent unwanted protein and DNA attachment to the surface. After thoroughly washing the chamber by flowing in excess volumes of experiment-specific reaction buffer (see Sections 3.2 and 4.3 for details), proper concentration of DNA and microspheres are sequentially flowed into the chamber to form DNA tethers. The detailed procedure is: (1) Flow in 1 volume (15 μL) of antidigoxigenin solution (20 ng/μL) in PBS and incubate for 5 min. (2) Wash with 5 volumes of blocking buffer (5 mg/mL casein sodium slat from bovine milk in reaction buffer) and incubate with residual blocker for 5 min. (3) Wash with 5 volumes of reaction buffer and sequentially flow in 1 volume of diluted DNA template in the sample buffer. Incubate for 10 min. (4) Wash with 5 volumes of sample buffer and sequentially flow in 1 volume of streptavidin-coated polystyrene microspheres (5 pM in blocking buffer). Incubate for 10 min. (5) Wash with 10 volumes of reaction buffer with one or more replicative proteins of interest and Mg2+ at proper concentrations and place the sample chamber on the optical setup for experiments. It is worth noting here that DNA samples need to be diluted to a proper concentration to achieve an optimal surface tether density under singlemolecule conditions. Low DNA concentrations will cause low tether density, making it difficult to spot a suitable tether in the chamber. Conversely, high DNA concentrations will lead to multiple DNA molecules attaching to one microsphere. The optimal concentration of DNA depends on the

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specific DNA template design and construction. In practice, tens to hundreds of pico molar of DNA sample is needed to achieve an acceptable tether density.

3. MOLECULAR MECHANISMS OF INDIVIDUAL PROTEINS IN THE REPLISOME REVEALED BY OPTICAL-TRAPPING TECHNIQUES 3.1 DNA Template Design and Construction Here, we detail the construction of DNA templates that can be used with optical-trapping system for the studies of individual proteins in the replisome (helicase or polymerase). A template generally consists of two segments: an anchoring segment and an unwinding segment, separated by a nick, which allows the DNA to be mechanically unwound (unzipped) (Fig. 3D) (Dechassa et al., 2011; Hall et al., 2009; Jiang et al., 2005; Jin et al., 2010;

Fig. 3 Unwinding template construction. A DNA template for the studies of individual proteins in the replisome is a product of ligation (D) of digoxigenin-labeled anchoring segment (A) and biotin-labeled unwinding segment. The unwinding segment consists of a biotin-labeled segment (B) and experiment-specific unwinding segment (C). See text for a detailed description of the experimental procedures.

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Koch & Wang, 2003; Koch et al., 2002; Li et al., 2015; Shundrovsky, Smith, Lis, Peterson, & Wang, 2006; Sun et al., 2011). The end of the anchoring segment is labeled by digoxigenin for attachment to the coverslip, and a biotin in the unwinding segment near the nick may be attached to a streptavidin-coated microsphere. By moving the coverslip away from the trapped microsphere to unzip the unwinding segment mechanically, a replication fork will be generated, allowing helicase loading/unwinding or polymerase synthesis. To provide sufficient separation between the anchor point and the unwinding segment while minimizing the Brownian motion of the trapped microsphere, the anchoring segment with an end-labeled digoxigenin tag is generally 1–2 kbp long. The unwinding segment is made of a biotin tagged segment followed by an experiment-specific target sequence. The length of the unwinding segment can vary from 800 to 4000 bp. As an example, we provide later a detailed protocol for constructing a 5.2-kbp DNA template containing a 1.1-kbp anchoring segment and a 4.1-kbp unwinding segment ( Johnson et al., 2007). The unwinding segment is derived from 17 pseudorepeats (or 17mer) of the 5S rRNA sequence, consisting of three five-repeats (or 5mer) of 207 bp, joined together by a 224 bp linking region. See later and Fig. 3 for detailed procedure. Anchoring segment preparation (Fig. 3A) (1) PCR amplify the anchoring segment from plasmid pRL574 (Koch et al., 2002). The forward primer (all primers and oligonucleotides were purchased from Integrated DNA Technologies and are listed in Table 1, unless specified otherwise) contains a 50 -digoxigenin label, designed to be 1.1 kbp away from the single BstXI (New England Biolabs, NEB) cutting site located on the plasmid. (2) BstXI digest the PCR product to generate a 30 -overhang for ligation of the unzipping segment. Unwinding segment preparation (3) Anneal two oligonucleotides to form a short biotin segment (Fig. 3B). One of the oligonucleotides is 50 -biotin labeled and lacks a phosphate for the generation of a nick. After annealing, two overhangs are generated for the ligation to the anchoring segment and 17mer segment. (4) The 17mer segment is a digestion product from plasmid pCP681 (Fig. 3C). After cutting with EarI (NEB), the desired segment is purified using an agarose gel purification kit (Zymo Research).

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Table 1 Primers and Oligonucleotides Used for DNA Templates Construction

5.2 kbp Anchoring nicked DNA segment template

Biotin segment

Forward primer

50 -/dig/ GTT GTA AAA CGA CGG CCA GTG AAT

Reverse primer

50 -CCG TGA TCC AGA TCG TTG GTG AAC

Biotin oligo

50 -/bio/ GAG CGG ATT ACT ATA CTA CAT TAG AAT TCG GAC

Complementary 50 -/phos/ GCT GTC TGA ATT CTA ATG TAG TAT oligo AGT AAT CCG CTC ATC G Y-shaped lesioncontaining DNA template

Arm 1

Arm 2

Adapters

Upstream segment

Forward primer

50 -/dig/ GTT GTA AAA CGA CGG CCA GTG AAT

Revise primer

50 -GAT CCA GAT CGT TGG TGA AC

Forward primer

50 -/bio/ GAT GCT TTT CTG TGA CTG GTG AG

Reverse primer

50 -ACG GTT ACC AGC CTA GCC GGG TCC TCA

Adapter 1

50 -/phos/ GCA GTA CCG AGC TCA TCC AAT TCT ACA TGC CGC

Adapter 2

50 -/phos/ GCC TTG CAC GTG ATT ACG AGA TAT CGA TGA TTG CGG CGG CAT GTA GAA TTG GAT GAG CTC GGT ACT GCA TCG

Adapter 3

50 -/phos/ GTA ACC TGT ACA GTG TAT AGA ATG ACG TAA CGC GCA ATC ATC GAT ATC TCG TAA TCA CGT GCA AGG CCT A

Forward primer

50 -CGC AGC TAC TGA GCC AGT CTG GTC ACA AGC G

Reverse primer

50 -GAT GGT CTC ACG GTT GGC GTC ATC GTG T Continued

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Table 1 Primers and Oligonucleotides Used for DNA Templates Construction—cont’d

Lesion segment

Lesion oligo

50 -/phos/ GGT GTC ACC AGC AGG CCG ATT GGG TT (CPD lesion) G GGT ATT CGC CGT GTC CCT CTC GAT GGC TGT AAG TAT CCT ATA GG

Complementary 50 -/phos/ ACC GCC TAT AGG ATA CTT ACA GCC oligo ATC GAG AGG GAC ACG GCG AAT ACC CAAC CCA ATC GGC CTG CTG GTG ACA CCC GAT Downstream Forward primer segment Revise primer

50 -TCA CCA ACG ATC TGG ATC ACG 50 -CGG TTG GCG TCA TCG TGT

(5) Ligate the 17mer segment with the biotin segment (1:10 molar ratio) at 16°C for 4 h using T4 ligase (NEB) to form the unwinding segment. Purify the ligated products using agarose gel purification to remove excess biotin segment. Anchoring segment and unwinding segment ligation (6) The final product is produced by ligating the anchoring segment and the unwinding segment using T4 ligase (in a 1:1 molar ratio) (Fig. 3D). Overnight ligation is normally necessary to maximize the ligation yield. A complete template is stable for a few days at 4°C. An advantage of this DNA template design is that, in the experiments, the DNA that has failed the final ligation step is automatically excluded, as the biotin for microsphere attachment is located on the unwinding segment such that no tether can be formed without this segment.

3.2 Helicase-Unwinding Assay Here, we detail the experimental procedures, using the aforementioned DNA template, to study T7 helicase unwinding (Fig. 4A) ( Johnson et al., 2007; Sun et al., 2011). T7 helicase consists of homologous monomers that form a ringshaped hexamer (Donmez & Patel, 2006). It binds one strand of dsDNA within its central channel, while excluding the complementary strand

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Fig. 4 Helicase unwinding and translocation assays. (A) Experimental configuration for helicase unwinding and translocations assays by a single T7 helicase. (B) and (C) correspond to helicase unwinding and translocation assays, respectively, in 2 mM dTTP. See text for a detailed description of the experimental procedures. Adapted from Johnson, D. S., Bai, L., Smith, B. Y., Patel, S. S., & Wang, M. D. (2007). Single-molecule studies reveal dynamics of DNA unwinding by the ring-shaped T7 helicase. Cell, 129, 1299–1309, with permission from the publisher.

(Ahnert & Patel, 1997). First, prepare T7 helicase by diluting helicase in its reaction buffer (50 mM NaCl, 3 mM EDTA, and 0.02% Tween-20 in 20 mM Tris–HCl, pH 7.5), followed by incubating 2 μM helicase, in the presence of 2 mM NTP and 3 mM EDTA, for 20 min to form a hexamer in which form T7 helicase is able to load onto ssDNA. Prior to flowing it into the sample chamber, the solution is diluted to a proper concentration (depending on the type of nucleotide used, see later), in 2 mM NTP, 3 mM EDTA, and 7 mM MgCl2 ( Johnson et al., 2007; Patel et al., 2011; Sun et al., 2011). A detailed procedure to detect helicase unwinding is provided as follows (Fig. 4B): (1) Mechanically unzipping to generate ssDNA. A tethered microsphere is used to stretch out the DNA under a “velocity clamp” mode, where the coverslip is moved away from the trapped microsphere at constant rate.

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The position of the microsphere in the trap is kept constant by modulating the light intensity (trap stiffness) of the trapping laser. In this mode, the dsDNA in the unwinding segment is mechanically unzipped to generate ssDNA. (2) Helicase loading. After 400 bp of dsDNA is mechanically unzipped, the setup is switched to a “hold” mode, in which the piezo stage and the light intensity are kept constant. Freely diffusing helicase can load onto the ssDNA, translocate from 50 to 30 , and subsequently unwind the remaining dsDNA, resulting in an increase of ssDNA length and, thereby, a decrease in its tension. (3) Helicase unwinding under constant force. The “hold” mode is exited to a “force clamp” mode after the force becomes lower than a preset value which is below that to unzip the DNA mechanically. In the “force clamp” mode, the force on the DNA tether is kept constant by adjusting both the position of the piezo stage and the light intensity. Therefore, the kinetics of helicase unwinding is measured by following the fork junction motion in real time. To ensure there is a single T7 helicase on the template, helicase concentration needs to be low enough so that the average helicase arrival time at the fork junction is significantly longer than the typical measurement time ( Johnson et al., 2007). In the presence of dTTP, we typically used 0.3 nM hexameric helicase (2 nM monomer) as the final concentration in our experiments. ATP binds to T7 helicase with a weaker affinity than dTTP (Hingorani & Patel, 1996; Lee & Richardson, 2010; Matson & Richardson, 1983), and thus a higher concentration of T7 helicase (e.g., 10 nM hexameric helicase) is recommended in the presence of ATP. Helicase slippage was also observed in this condition (see Section 6.1). For the experiments in which both nucleotides are present, 1 nM hexameric helicase is an appropriate concentration to achieve single helicase activity.

3.3 Helicase Translocation Assay The helicase translocation assay aims to investigate the ssDNA translocation rate of helicase on long stretches of ssDNA ( Johnson et al., 2007). We have used the unzipping fork to mark the initial and final positions and times of the helicase translocation. The helicase’s arrival at a fork and its subsequent unwinding of the fork leads to a force drop, which marks the initial position and start time of the helicase translocation. Immediately following this detection, a region of downstream ssDNA is rapidly generated via

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mechanically unzipping. Helicase then translocates on this newly available ssDNA until it catches up with, and unwinds, the downstream fork, leading to another force drop which marks the final position and time for the helicase translocation. The ssDNA translocation rate is obtained from the distance that the helicase travels over the time the helicase takes to translocate this distance. The detailed procedure is listed below (Fig. 4C): (1) Mechanically unzipping to generate ssDNA. Using a “velocity clamp” mode, dsDNA in the unwinding segment is mechanically unzipped to provide a segment of ssDNA. (2) Detection of helicase loading. The setup is switched to a “hold” mode, and the DNA extension is maintained until the force drops below a threshold indicating helicase unwinding of the DNA fork. (3) Mechanically unzipping to generate more ssDNA. Mechanically unwind about 600 bp to generate a new ssDNA region for helicase translocation. Transition to a “hold” mode again. (4) Detection of helicase arrival at the fork. Maintain the new DNA fork position until force drops again, indicating that the helicase has caught up with the fork. Using this method, it should be noted that the translocation rate on ssDNA is obtained under a tension of 14 pN. Around this force, the translocation rate of T7 helicase does not show a strong dependence on force on the ssDNA ( Johnson et al., 2007).

3.4 Polymerase Strand Displacement Assay Replicative DNAPs are responsible for faithfully synthesizing genomic DNA, in both prokaryotes and eukaryotes, by adding nucleotides to the 30 terminus of the primer strand. To increase the fidelity of DNA replication, replicative DNAPs typically have a 30 –50 exonuclease to allow for proofreading by excision of erroneous incorporated nucleotides. DNAP’s activity can be measured with a strand displacement assay in which DNAP carries leading-strand synthesis while displacing the lagging strand at a DNA fork. As the DNA fork is typically an obstacle for DNAP, a force assisting strand separation will facilitate the advancement of DNAPs (Fig. 5A). Here, using T7 DNAP as an example, we detail the experimental procedures of the strand displacement assay on the aforementioned DNA template (Sun et al., 2015). Wild-type (wt) DNAP (NEB) contains gp5 and trx and can be used directly. Exo DNAP is assembled by adding 10 μM of gp5 to 50 μM E. coli trx and incubating at room temperature for 5 min. Before data

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Fig. 5 Polymerase strand displacement assay. (A) Experimental configuration for the polymerase strand displacement assay. This cartoon illustrates the experimental configuration for the observation of DNA synthesis by a single T7 DNAP as it displaces the other strand. (B) A representative trace showing the number of replicated base pairs vs time in the presence of 1 mM dNTPs under 12 pN.

acquisition, the appropriate DNAP is diluted in its reaction buffer (50 mM Tris–HCl pH 7.5, 40 mM NaCl, 10% glycerol, 1.5 mM EDTA, 2 mM DTT, 8 mM MgCl2, and 1 mM dNTP each) to 30 nM, before flowing it into the chamber. The experimental procedure is similar to the helicaseunwinding assay: (1) Mechanically unzipping to generate ssDNA. At a constant velocity, several hundred base pairs of dsDNA are mechanically unzipped to produce a region of ssDNA as a template for the polymerase. (2) Polymerase loading. DNA length is maintained under a “hold” mode until a force drop is observed, indicating polymerase unwinding of the DNA fork. (3) Polymerase unwinding/synthesizing under constant force. The setup then switches to a “force clamp” mode to maintain a constant force, while a polymerase unwinds and synthesizes the downstream dsDNA. It is worth noting here that wt T7 DNAP has force-dependent synthesis and exonuclease activities. Under a force less than 8 pN, exonuclease activity dominates under the influence of the reannealing fork, and this leads to a decrease in DNA extension corresponding to a decrease in the number of base pairs replicated (see later). At higher forces, this effect is significantly alleviated, and thus wt T7 DNAP can perform strand displacement synthesis (Fig. 5B). The experimental configuration and DNA template design detailed ealier provide several advantages for detecting the motion of helicase and polymerase. First, the motor protein of interest does not need to be tagged

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or anchored, nor under direct mechanical stress, minimizing potential perturbation to the protein. Second, for proteins that can only associate with ssDNA, experiments can only be initiated after a ssDNA loading region is mechanically generated. Thus, the start of an experiment on each DNA molecule can be independently controlled. Third, for both helicase unwinding and polymerase synthesis assays, each base pairs of dsDNA unwound results in the release of 2 nt ssDNA (for helicase) or 1 nt ssDNA and 1 bp dsDNA (for polymerase), amplifying the detection signal. Using this method, we have directly measured the dynamic activities of individual proteins in the replisome ( Johnson et al., 2007; Sun et al., 2011, 2015). Next, we expand this approach to investigate multicomponent, dynamic machineries of the T7 replisome, namely, more than one protein is investigated at any one time during replication.

4. SINGLE-MOLECULE STUDIES OF THE RESPONSE OF A REPLISOME TO DNA DAMAGE DNA replication often relies on DNA damage tolerance pathways to overcome DNA damage which, if not repaired, may cause a replication fork to stall or collapse, leading to genomic instability and cell death. In order to complete the cell cycle and maintain cell survival, it is often more advantageous to circumvent blocked replication forks and postpone the damage repair (Yeeles, Poli, Marians, & Pasero, 2013). Thus, a detailed understanding of the response of a replisome to DNA damage, as well as possible lesion tolerance pathways, is of great interest to multiple fields, including genome stability, cell survival, and human disease (Zeman & Cimprich, 2014). Previous studies showed that the translocation of T7 helicase on ssDNA was blocked by bulky DNA adducts (Brown & Romano, 1989). Therefore, damage to DNA could interfere with the helicase in its translocation or unwinding, leading to the stalled or collapsed replication fork. T7 DNAP belongs to the high-fidelity Pol A family of polymerases and is unable to bypass UV-induced lesions on its own. However, the exonuclease-deficient (exo) T7 DNAP mutant is able to bypass them, suggesting its proofreading activity is important in its ability to bypass a lesion (McCulloch & Kunkel, 2006; Smith, Baeten, & Taylor, 1998). These processes are highly dynamic, which is often averaged out in the classical biochemical experiments. Thus, we have developed single-molecule methods to investigate these questions (Sun et al., 2015).

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4.1 DNA Template Design and Construction To monitor the real-time dynamic process of DNA replication and investigate the response of each replicative protein after encountering a lesion, we designed a Y-shaped DNA template containing a single cis–syn cyclobutane pyrimidine dimer (CPD) lesion in either the leading-strand or laggingstrand of a forked DNA template (Inman et al., 2014; Sun et al., 2015). Although we use CPD as an example, the methods described here could be easily applied to other types of lesions. This DNA template consists of three distinct dsDNA segments: two arms and a trunk (Fig. 6). The two arms can be amplified from plasmids with end-labeled tags, followed by restriction enzyme cuts to create overhangs for subsequent ligation. The trunk is, itself, a ligation product of a three-piece segment: upstream segment, lesion segment, and downstream segment. The upstream and downstream segments are made via PCR amplification of plasmids, and ligation overhangs are generated by restriction enzyme digest. The two arms and the trunk are linked via three adapter oligonucleotides, and upon ligation, form Y-shaped DNA template. The resulting template provides an artificial replication fork containing a ssDNA loading region for the replisome to load and replicate. Here, we provide a detailed procedure to construct this lesion-containing DNA template (Fig. 6): Arms preparation (Fig. 6A) (1) Arm 1 (1.1 kbp) is amplified from plasmid pLB574 using a digoxigeninlabeled primer. Arm 2 (2 kbp) is amplified from plasmid pBR322 (NEB) using a biotin-labeled primer. (2) Arm 1 and Arm 2 PCR products are digested with BstXI (NEB) and BstEII (NEB), respectively, to create overhangs for adapter ligation. (3) Arm 1 is annealed to a short DNA at 1:10 molar ratio at 16°C for at least 4 h. The short DNA is an annealing product of two adapters (adapters 1 and 2) with an overhang complementary to that of Arm 1. Ligation products are purified using agarose gel purification kit (Zymo Research). (4) Arm 2 is annealed to adapter 3 at 1:10 molar ratio, and gel purified to eliminate excess adapters. (5) Adapter 2 from Arm 1 (step 3) and adapter 3 from Arm 2 (step 4) are partially complementary to each other and are annealed to create a replication fork with a short 30-bp trunk with 3-bp overhang for subsequent ligation to the trunk segment.

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Fig. 6 Lesion-containing template construction. A DNA template for studies of the response of a replisome to a DNA lesion is illustrated. Two arms are linked via three adapters (A). The trunk consists of an upstream segment (B), a lesion-containing segment (C), and a downstream segment (D). The final DNA template with a lesion located on the leading strand (E) or the lagging strand (F) is a ligation product of two arms and a trunk. See text for a detailed description of the procedures.

Trunk preparation (6) For the trunk, a 1.1-kbp upstream (Fig. 6B) and 1.1-kbp downstream segment (Fig. 6D) are amplified via PCR from a pRL574 plasmid variant.

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(7) The upstream and downstream PCR products are then cut by BsaI and BstXI (NEB), respectively, to create overhangs for ligation with the lesion segment. (8) The lesion segment (Fig. 6C) is made by annealing a CPD lesioncontaining oligonucleotide with its complementary oligonucleotide. Of note, the cis–syn cyclobutane thymine dimer phosphoramidite is purchased from Glen Research (Sterling, VA) and was used for synthesis of the CPD oligonucleotide with PAGE purification by Oligos Etc (Wilsonville, OR). The upstream segment, the lesion segment, and the downstream segment are ligated and purified using gel purification. Arms and trunk ligation (9) To create a DNA template with a CPD lesion located on the leadingstrand template, the upstream DNA segment is also digested with AlwNI (NEB) before ligation with the lesion segment. This creates an overhang for ligation with arms and results in a CPD lesion located at 1145–1146 bp from the initial fork (Fig. 6E). To create the DNA template with a CPD lesion located on the lagging-strand template, the trunk sequence was flipped, and the downstream DNA segment, instead of the upstream DNA segment, is digested with AlwNI (NEB) to create an overhang for ligation with the arms, resulting in a CPD lesion located at 1223–1224 bp from the initial fork (Fig. 6F). Arms are ligated with the trunk at 1:4 molar ratio under 16°C for 3 h on the day of an experiment.

4.2 Response of Individual Proteins to DNA Damage: HelicaseUnwinding Assay and Polymerase Strand Displacement Assay. To investigate the response of the bacteriophage T7 replisome to a DNA lesion, the effect of the lesion on individual proteins need to be first examined (Fig. 7) (Sun et al., 2015). Helicase unwinding and polymerase strand displacement experiments are conducted as previously described in Sections 3.2 and 3.4. In brief, several hundred base pairs of dsDNA before the lesion are mechanically unzipped (with an average unzipping force of 15 pN), at a constant velocity of 1400 nm/s, to produce a ssDNA loading region for helicase or a template for polymerase. Then, DNA length is maintained until a force drop is observed (below a predetermined threshold), indicating helicase or polymerase unwinding of the DNA fork. Subsequently, a constant force is maintained, while helicase or polymerase

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Fig. 7 Helicase unwinding and polymerase synthesizing through a cis–syn CPD lesion. (A) Helicase unwinding through a CPD lesion. This cartoon illustrates the experimental configuration for the observation of T7 helicase unwinding through a CPD lesion. A CPD lesion (red star) is located in the leading-strand DNA. Representative traces show the number of unwound base pairs vs time in the presence of 2 mM dTTP. (B) Polymerase synthesizing through a CDP lesion. The cartoon illustrates the experimental configuration for the observation of T7 DNAP synthesizing through a CPD lesion. Representative traces show the number of replicated base pairs vs time in the presence of 1 mM dNTPs under 12, 8, and 6 pN. Note that at 6 pN, DNAP excised DNA from the 30 end. The dotted lines indicate the lesion position. Adapted from Sun, B., Pandey, M., Inman, J. T., Yang, Y., Kashlev, M., Patel, S. S., et al. (2015). T7 replisome directly overcomes DNA damage. Nature Communications, 6, 10260, with permission from the publisher.

unwinds the dsDNA. To be consistent, both experiments are conducted in a replication buffer which consists of 50 mM Tris–HCl (pH 7.5), 40 mM NaCl, 10% glycerol, 1.5 mM EDTA, 2 mM DTT and 1 mM dNTPs (each), and 8 mM MgCl2. For the helicase-unwinding assay, 0.4 nM hexamer helicase is typically used to ensure single helicase conditions. For the strand displacement assay, 30 nM of the appropriate DNAP in 50 μL replication buffer is flowed into the chamber before data acquisition.

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4.3 Response of a Replisome to DNA Damage: Leading-Strand Replication Assay The leading-strand replication assay is performed under a “force clamp” (Fig. 8A) (Sun et al., 2015). To avoid strand displacement synthesis by DNAP alone, the experiments are conducted under a low force of 6 pN where T7 DNAP exonuclease activity alone would lead to a decrease in DNA length. Therefore, the DNA length increase would only be due to either helicase unwinding or helicase/DNAP replication under this condition. These two scenarios are differentiated by the rate of the DNA length increase which is utilized to determine the fates of the proteins after the lesion (see Section 5.4 for details). The helicase and polymerase are prepared as follows: first, 30 nM of the appropriate helicase hexamer is incubated for 10 min, on ice, in the replication buffer, then 30 nM of the appropriate DNAP is added, and the solution is incubated for 10 min at room temperature. The resulting 50 μL solution is flowed into a chamber just before data acquisition. The DNA template design contains a 27-nt initial ssDNA region (Fig. 6E) that accommodates only one helicase with one DNAP loaded at the fork, as each T7 helicase has been shown to bind and protect 25–30 bases of ssDNA (Egelman, Yu, Wild, Hingorani, & Patel, 1995; Hingorani & Patel, 1993; Patel & Hingorani, 1993). The low concentrations of helicase and DNAP, added at a stoichiometric ratio, ensure that the experiments are likely to occur under a single copy conditions.

5. DATA ANALYSIS After the acquired data signals are converted into force and DNA extension, the data need to be further processed as described below.

5.1 DNA Elastic Parameter Determination Elasticity parameters, of both dsDNA and ssDNA, are necessary for data conversion and are obtained from the DNA force–extension measurements. They are strongly dependent on the buffer conditions used in the experiments. The force–extension relation of dsDNA was obtained by stretching dsDNA in the same replication buffer mentioned earlier and fit by using a modified Marko–Siggia worm-like-chain model (Wang, Yin, Landick, Gelles, & Block, 1997). The fit yielded the following fitting parameter values (Sun et al., 2015): the contour length per base is 0.338 nm, the persistence length of DNA is 44.5 nm, and the stretch modulus is 1200 pN.

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Fig. 8 Leading-strand synthesis on a DNA template containing a CPD lesion in the presence of helicase. (A) Experimental configuration. This cartoon illustrates the experimental configuration for the observation of leading-strand synthesis through a CPD lesion (red star). (B) Representative traces showing DNA length vs time for a wt DNAP with helicase in the presence of 0.5 mM dNTPs and 6 pN. The dotted lines indicate the position of a single CPD lesion. For clarity, traces have been shifted along the time axis. Cartoons illustrate different protein compositions at the fork before and after the lesion for each trace. Insets display the distributions of DNA length increase rates before and after the lesion. Adapted from Sun, B., Pandey, M., Inman, J. T., Yang, Y., Kashlev, M., Patel, S. S., et al. (2015). T7 replisome directly overcomes DNA damage. Nature Communications, 6, 10260, with permission from the publisher.

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The force–extension relation of ssDNA may be described by an extensible freely jointed chain (FJC) model (Smith, Cui, & Bustamante, 1996). For forces higher than 12 pN, this relation may be obtained using an unzipping template with a hairpin at its distal end ( Johnson et al., 2007). Upon mechanically unzipping all the way to the hairpin to generate ssDNA, the ssDNA may then be stretched to obtain the force–extension curve of the ssDNA. The resulting force extension is well fit by the FJC model, yielding a contour length per base of 0.52 nm, a Kuhn’s length of 1.91 nm and a stretch modulus of 393 pN. For forces lower than 12 pN, the force extension is directly determined using a helicase-based method ( Johnson et al., 2007). In brief, the helicase loads onto the ssDNA and starts to unwind dsDNA under a constant force. This is followed by stretching the two ends of the DNA, at 1400 nm/s to 13 pN, which is sufficient to remove any secondary structures in the ssDNA. The force increase is rapid so that the helicase is not expected to move forward by more than a few base pair during this time. Thus, the force–extension measurements could then be used to determine the number of ssDNA base pairs at the force prior to stretching. Setting different constant low forces for the initial dsDNA unwinding by the helicase allows the determination of the force–extension curve of ssDNA over the entire range of relevant forces. The resulting curve may be fit with a polynomial ( Johnson et al., 2007).

5.2 DNA Extension Conversion to Number of Base Pairs For the helicase-unwinding studies, one base pair unwound generates two nucleotides of ssDNA. Accordingly, real-time DNA extension under a given force is converted into the number of base pairs unwound ( Johnson et al., 2007). To improve positional accuracy and precision, the data during the initial mechanically unzipping are used to align to a theoretical unzipping curve (Dechassa et al., 2011; Deufel & Wang, 2006; Hall et al., 2009; Inman et al., 2014; Jiang et al., 2005; Jin et al., 2010; Johnson et al., 2007; Li & Wang, 2012; Li et al., 2015; Shundrovsky et al., 2006). For the DNAP strand displacement synthesis studies, one separated base pair is converted into one base pair of dsDNA, via DNA synthesis, and one nucleotide of ssDNA. Accordingly, DNA extension under a given force is converted into the number of nucleotides synthesized, or excised, by DNAP (Sun et al., 2015). In particular, under 6 pN of force, a 1 nm increase in length corresponded to 1.95 bp unwound by helicase and 1.74 bp replicated by the leading-strand synthesis.

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To determine the expected lesion position in the extension signal on a lesion-containing template in the leading-strand replication assay, the DNA template before the lesion is assumed to be replicated, and the lesion position in base pairs which is known from the DNA template design is converted into extension (in nanometer).

5.3 Helicase Slippage Determination and Processivity Measurements In the helicase-unwinding assay, when ATP is used, a remarkable sawtooth pattern in the unwinding trace is observed as processive unwinding is interrupted by slippage events. The slippage events are due to helicase losing its grip on the ssDNA, and sliding backwards under the influence of the reannealing DNA fork (Sun et al., 2011). We set a threshold of 2000 bp/s, in the reverse velocity, for identifying slippage. Distances that a helicase travels between slips are compiled to determine processivity (Fig. 9A), and the distances follow an exponential distribution, indicating a stochastic process in slippage.

5.4 Unwinding Rate and Replication Rate Measurements and Comparison For the sequence-dependent helicase-unwinding studies, to improve positional accuracy and precision to a few base pairs, the initial mechanical unzipping section of each trace was aligned with the theoretical prediction ( Johnson et al., 2007; Sun et al., 2011). After alignment, an instantaneous unwinding rate at each sequence position is determined using a Gaussian weighted filter of 0.05 s and then resampling at 1-bp intervals along the sequence position. The final rate vs position curve is found by averaging each position over all of the traces. For the leading-strand replication studies, one first has to determine whether the extension change is due to the helicase alone or DNAP synthesis coupled with helicase unwinding. The differentiation is achieved by directly measuring and comparing the length increase rates in nanometer per second on an unmodified DNA template (Sun et al., 2015). The average rates are found from linear fits to each trace, followed by averaging over all of the traces. Taking the experiments using 0.5 mM dNTP (each) and 6 pN as an example, the unwinding by helicase alone results in a DNA length increase at rate of 63  22 nm/s, and when DNAP is present together with helicase, the rate increases to 111  13 nm/s (Sun et al., 2015). Subsequently, depending on whether the DNA template is replicated, the rates

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Fig. 9 Helicase slippage and a proposed coordinated model. (A) An example of unwinding with ATP to illustrate the method of determining distance between slips. (B) Hexameric helicase subunit coordination. Each subunit is uniquely labeled with a different color and has a potential ssDNA-binding site (small dots). Nucleotide binding and subsequent hydrolysis occur sequentially around the ring. If a subunit is nucleotide ligated (the state of hydrolysis indicated by Ni), it has a nonzero probability of being bound to ssDNA. During unwinding, the leading subunit can bind to a nucleotide (N) and thus acquire affinity for the downstream ssDNA. This stimulates the last nucleotide-bound subunit to release its nucleotide and ssDNA. Then, the cycle proceeds again around the ring. Slippage occurs when all subunits simultaneously release ssDNA. Adapted from Sun, B., Johnson, D. S., Patel, G., Smith, B. Y., Pandey, M., Patel, S. S., et al. (2011). ATP-induced helicase slippage reveals highly coordinated subunits. Nature, 478, 132–135, with permission from the publisher.

in nanometer per second can be converted into base pairs per second using the elastic parameters of both ssDNA and dsDNA.

5.5 Fate of a Replisome Encountering DNA Damage As mentioned earlier, to examine the fate of a replisome encountering DNA damage, one has to determine whether the postlesion movement of the fork is due to the helicase alone or DNAP synthesis coupled with helicase unwinding. The length increase rate after the lesion is measured by a linear fit to the unwinding trace and then compared with the rates obtained from the unmodified template (Sun et al., 2015). DNAP may terminate synthesizing after the lesion but still associate with the helicase and alter its unwinding rate. To mimic this condition, the control experiments to measure helicase-unwinding rates are conducted in the presence of

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DNAP by using a modified template with a 30 inverted dT incorporated at the adapter 1 (IDT) from which DNAP could not synthesize (Sun et al., 2015).

6. UNIQUE FEATURES OF THE BACTERIOPHAGE T7 REPLISOME REVEALED BY SINGLE-MOLECULE OPTICAL-TRAPPING TECHNIQUES Here, we summarize major results obtained from the studies of the bacteriophage T7 replisome using single-molecule optical-trapping techniques. For more specific details regarding these experiments, or data analysis, we refer the reader to the original publications (Sun et al., 2011, 2015).

6.1 Helicase Slippage and Subunit Coordination T7 helicase is a model hexameric helicase that uses dTTP to unwind dsDNA. Earlier bulk studies of T7 helicase found that although T7 helicase is capable of ATP hydrolysis, it did not unwind DNA efficiently in the presence of ATP (Hingorani & Patel, 1996; Matson & Richardson, 1983). Our single-molecule results revealed that ATP supported not only dsDNA unwinding but also a significantly faster unwinding rate than that with dTTP (Sun et al., 2011). However, in the presence of ATP, helicase unwinding is frequently interrupted by slippage, where helicase loses the grip of ssDNA, moves in a reverse direction along the ssDNA, but then regains the grip of ssDNA before moving forward again. These results have resolved the mystery of the apparent lack of significant unwinding activity seen in bulk studies, as the frequent slippage prevents helicase from moving over a substantial distance to be detected in a strand separation assay. Thus far, T7 helicase is the only motor protein that has reported nucleotide-specific slippage behavior. Slippage has been observed with other helicases, however, it appears to result from different causes (Klaue et al., 2013; Lee et al., 2014; Manosas, Spiering, Ding, Croquette, & Benkovic, 2012; Myong, Bruno, Pyle, & Ha, 2007; Myong, Rasnik, Joo, Lohman, & Ha, 2005; Qi, Pugh, Spies, & Chemla, 2013; Sun et al., 2008). More importantly, these slippage events provided us with a unique opportunity to investigate how different subunits of the helicase coordinate their mechanical and chemical activities (Sun et al., 2011). By examining the helicase processivity in a mixture of ATP and dTTP in conjunction with theoretical modeling, we found that all, or nearly all, six of subunits of the helicase must coordinate their mechanochemical activities. Based on

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structural studies of hexameric ring-shaped helicases E1 and Rho (Enemark & Joshua-Tor, 2006; Thomsen & Berger, 2009), we proposed a mechanistic model for T7 helicase. In this model, coordination could occur sequentially around the hexameric ring with the leading subunit poised for NTP binding and each successive subunit having a bound nucleotide in states of progression along the chemical reaction pathway. Once the leading subunit binds to an NTP and reels in the DNA, the remaining subunits progress to their next reaction states. Product release by the last participating subunit results in release of DNA from that subunit, and thus completes a single cycle (Fig. 9B). It is speculated that slippage may provide an evolutionary advantage for replication and allow helicase synchronization with a nonsynthesizing or slow-moving DNAP.

6.2 Inherent Tolerance of the T7 Replisome to DNA Damage Leading-strand DNA lesions are often major obstacles for replication progression, as the high-fidelity replicative polymerase is incapable of directly proceeding through them. As examples, for both T4 and E. coli replisomes, leading- and lagging-strand DNA replication becomes uncoupled after encountering a leading-strand DNA lesion (Higuchi et al., 2003; McInerney & O’Donnell, 2007; Nelson & Benkovic, 2010). A replisome often adopts an indirect pathway to tolerate DNA damage, which requires the replisome to avoid lesions and/or reinitiate replication after bypass (Yeeles & Marians, 2011; Yeeles et al., 2013). Using the techniques mentioned earlier, we examined how T7 helicase and polymerase deal with a UV-induced CPD lesion in the leading-strand template (Sun et al., 2015). We demonstrated that, in the presence of T7 helicase, a substantial fraction of T7 DNAP is able to replicate through the lesion directly. The helicase and the polymerase replicate up to the lesion and stay together to synthesize through the lesion through specific helicase–DNAP interactions. Upon lesion bypass, DNAP and the helicase concurrently resume their independent activities (Fig. 8C). These results suggest that the T7 replisome is fundamentally permissive of DNA lesions. This is, to our knowledge, the first observation of CPD tolerance by a helicase-coupled replicative polymerase synthesizing through a lesion, rather than circumventing it. Based on these results, we proposed a new lesion bypass pathway, in which a replicative DNAP directly synthesizes through a leading-strand lesion with the assistance of a helicase. In contrast to other pathways, the new pathway functions in the absence of additional accessory proteins, with the exception of

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helicase, and replication fork adjustment and reinitiation of the replisome are not required. Therefore, it is more efficient in terms of replisome recovery.

7. CONCLUSIONS Single-molecule optical-trapping techniques have expanded to impact a wide field of biological sciences. The experimental methods detailed here offer the ability to resolving helicase or polymerase motion at high spatial and temporal resolution, generating a more comprehensive understanding of these motor proteins and their corresponding functions in the replisome during replication. The detailed experimental design mentioned here provides an opportunity to monitor the dynamics of the multicomponent machinery of a replisome and is capable of elucidating the cooperation and coordination of two or more proteins simultaneously. We anticipate that this method will continue to play an important role in the study of proteins and molecular machines and will be further enhanced in combination with other single-molecule techniques.

ACKNOWLEDGMENTS We thank Dr. Shanna M. Moore from the Wang Laboratory at Cornell University for critical comments on the manuscript. We wish to acknowledge support from Shanghai Pujiang Program (16PJ1406900 to B.S.), National Key Research and Development Program of China (2016YFA0500900 to B.S.) and National Science Foundation Grant (MCB1517764 to M.D.W.).

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Li, M., & Wang, M. D. (2012). Unzipping single DNA molecules to study nucleosome structure and dynamics. Methods in Enzymology, 513, 29–58. Manosas, M., Spiering, M. M., Ding, F., Croquette, V., & Benkovic, S. J. (2012). Collaborative coupling between polymerase and helicase for leading-strand synthesis. Nucleic Acids Research, 40, 6187–6198. Matson, S. W., & Richardson, C. C. (1983). DNA-dependent nucleoside 50 -triphosphatase activity of the gene 4 protein of bacteriophage T7. The Journal of Biological Chemistry, 258, 14009–14016. Matson, S. W., Tabor, S., & Richardson, C. C. (1983). The gene 4 protein of bacteriophage T7. Characterization of helicase activity. The Journal of Biological Chemistry, 258, 14017–14024. McCulloch, S. D., & Kunkel, T. A. (2006). Multiple solutions to inefficient lesion bypass by T7 DNA polymerase. DNA Repair, 5, 1373–1383. McInerney, P., & O’Donnell, M. (2007). Replisome fate upon encountering a leading strand block and clearance from DNA by recombination proteins. The Journal of Biological Chemistry, 282, 25903–25916. Moffitt, J. R., Chemla, Y. R., Smith, S. B., & Bustamante, C. (2008). Recent advances in optical tweezers. Annual Review of Biochemistry, 77, 205–228. Myong, S., Bruno, M. M., Pyle, A. M., & Ha, T. (2007). Spring-loaded mechanism of DNA unwinding by hepatitis C virus NS3 helicase. Science, 317, 513–516. Myong, S., Rasnik, I., Joo, C., Lohman, T. M., & Ha, T. (2005). Repetitive shuttling of a motor protein on DNA. Nature, 437, 1321–1325. Nelson, S. W., & Benkovic, S. J. (2010). Response of the bacteriophage T4 replisome to noncoding lesions and regression of a stalled replication fork. Journal of Molecular Biology, 401, 743–756. Patel, S. S., & Hingorani, M. M. (1993). Oligomeric structure of bacteriophage T7 DNA primase/helicase proteins. The Journal of Biological Chemistry, 268, 10668–10675. Patel, G., Johnson, D. S., Sun, B., Pandey, M., Yu, X., Egelman, E. H., et al. (2011). A257T linker region mutant of T7 helicase-primase protein is defective in DNA loading and rescued by T7 DNA polymerase. The Journal of Biological Chemistry, 286, 20490–20499. Qi, Z., Pugh, R. A., Spies, M., & Chemla, Y. R. (2013). Sequence-dependent base pair stepping dynamics in XPD helicase unwinding. eLife, 2, e00334. Shundrovsky, A., Smith, C. L., Lis, J. T., Peterson, C. L., & Wang, M. D. (2006). Probing SWI/SNF remodeling of the nucleosome by unzipping single DNA molecules. Nature Structural & Molecular Biology, 13, 549–554. Singleton, M. R., Dillingham, M. S., & Wigley, D. B. (2007). Structure and mechanism of helicases and nucleic acid translocases. Annual Review of Biochemistry, 76, 23–50. Smith, C. A., Baeten, J., & Taylor, J. S. (1998). The ability of a variety of polymerases to synthesize past site-specific cis-syn, trans-syn-II, (6–4), and Dewar photoproducts of thymidylyl-(30 –>50 )-thymidine. Journal of Biological Chemistry, 273, 21933–21940. Smith, S. B., Cui, Y., & Bustamante, C. (1996). Overstretching B-DNA: The elastic response of individual double-stranded and single-stranded DNA molecules. Science, 271, 795–799. Sun, B., Johnson, D. S., Patel, G., Smith, B. Y., Pandey, M., Patel, S. S., et al. (2011). ATPinduced helicase slippage reveals highly coordinated subunits. Nature, 478, 132–135. Sun, B., Pandey, M., Inman, J. T., Yang, Y., Kashlev, M., Patel, S. S., et al. (2015). T7 replisome directly overcomes DNA damage. Nature Communications, 6, 10260. Sun, B., & Wang, M. D. (2016). Single-molecule perspectives on helicase mechanisms and functions. Critical Reviews in Biochemistry and Molecular Biology, 51, 15–25. Sun, B., Wei, K. J., Zhang, B., Zhang, X. H., Dou, S. X., Li, M., et al. (2008). Impediment of E. coli UvrD by DNA-destabilizing force reveals a strained-inchworm mechanism of DNA unwinding. The EMBO Journal, 27, 3279–3287.

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Tabor, S., Huber, H. E., & Richardson, C. C. (1987). Escherichia coli thioredoxin confers processivity on the DNA polymerase activity of the gene 5 protein of bacteriophage T7. The Journal of Biological Chemistry, 262, 16212–16223. Thomsen, N. D., & Berger, J. M. (2009). Running in reverse: The structural basis for translocation polarity in hexameric helicases. Cell, 139, 523–534. Wang, M. D., Yin, H., Landick, R., Gelles, J., & Block, S. M. (1997). Stretching DNA with optical tweezers. Biophysical Journal, 72, 1335–1346. Yeeles, J. T. P., & Marians, K. J. (2011). The Escherichia coli replisome is inherently DNA damage tolerant. Science, 334, 235–238. Yeeles, J. T. P., Poli, J., Marians, K. J., & Pasero, P. (2013). Rescuing stalled or damaged replication forks. Cold Spring Harbor Perspectives in Biology, 5, 1–15. Zeman, M. K., & Cimprich, K. A. (2014). Causes and consequences of replication stress. Nature Cell Biology, 16, 2–9.

CHAPTER FOUR

Recent Advances in Biological Single-Molecule Applications of Optical Tweezers and Fluorescence Microscopy M. Hashemi Shabestari*,1, A.E.C. Meijering*,1, W.H. Roos†, G.J.L. Wuite*, E.J.G. Peterman*,2 *Vrije Universiteit, Amsterdam, The Netherlands † Moleculaire Biofysica, Zernike Institute, Rijksuniversiteit Groningen, Groningen, The Netherlands 2 Corresponding author: e-mail address: [email protected]

Contents 1. Introduction 2. Instrumentation 3. Applications 3.1 Single Optical Trap 3.2 Dual-Trap Optical Tweezers 3.3 FRET Studies With Confocal Fluorescence Microscopy 4. Experimental Protocol 4.1 Experimental Setup 4.2 Methods 5. Conclusion Acknowledgments References

86 88 95 95 99 109 111 112 112 113 114 114

Abstract Over the past two decades, single-molecule techniques have evolved into robust tools to study many fundamental biological processes. The combination of optical tweezers with fluorescence microscopy and microfluidics provides a powerful single-molecule manipulation and visualization technique that has found widespread application in biology. In this combined approach, the spatial (
nm) and temporal (
ms) resolution, as well as the force scale (
pN) accessible to optical tweezers is complemented with the power of fluorescence microscopy. Thereby, it provides information on the local presence, identity, spatial dynamics, and conformational dynamics of single biomolecules. Together, these techniques allow comprehensive studies of, among others, molecular

1

These authors contributed equally.

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motors, protein–protein and protein–DNA interactions, biomolecular conformational changes, and mechanotransduction pathways. In this chapter, recent applications of fluorescence microscopy in combination with optical trapping are discussed. After an introductory section, we provide a description of instrumentation together with the current capabilities and limitations of the approaches. Next we summarize recent studies that applied this combination of techniques in biological systems and highlight some representative biological assays to mark the exquisite opportunities that optical tweezers combined with fluorescence microscopy provide.

1. INTRODUCTION The exploration of single-molecule systems and their interactions with each other in terms of mechanical forces has substantially benefitted from recent advances in the now well-established field of optical tweezers. The very high sensitivity (pN, nm, kBT) and the wide temporal resolution (s to ms) of optical tweezers can be used to measure noninvasively the mechanics of single molecules and their interactions with other molecules. It has been about three decades since Ashkin presented the first application of optical tweezers in biology (Ashkin, Dziedzic, & Yamane, 1987), and since then, optical tweezers have become one of the most widely used single-molecule tools in biology. What makes the use of optical tweezers even more compelling is its compatibility with various types of light microscopy, such as bright field, differential interference contrast, phase contrast, and fluorescence microscopy. Not surprisingly, optical trapping was soon combined with fluorescence microscopy as a method for visualizing single molecules. In the first combination study, a long piece of DNA was labeled with ethidium bromide and stretched between two beads held in two optical traps (Chu, 1991). The relaxation of the DNA upon release of one bead could be followed by simply watching the movements of the fluorescently labeled DNA. In the last two decades, a variety of fluorescence techniques have been successfully combined with optical tweezers. Epi-illuminated wide-field and total internal reflection fluorescence (TIRF) microscopy, being the most straightforward to implement, have been adopted first (Arai et al., 1999; Bianco et al., 2001; Funatsu et al., 1997; Saito, Aoki, & Yanagida, 1994; Sarangapani et al., 2014). More recently, confocal fluorescence microscopy and stimulated emission depletion (STED) super-resolution microscopy have been integrated with optical tweezers (Bornschl€ ogl, Romero, Vestergaard, & Joanny, 2013;

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Comstock et al., 2015; Duesterberg et al., 2015; Grashoff et al., 2010; Heller et al., 2013; Hohng et al., 2007; Wolfson et al., 2015; Zhou et al., 2011). Each of these visualization approaches has its own set of strengths and weaknesses for the analysis of protein dynamics. Addition of fluorescence microscopy enables direct visualization of individual proteins on biomolecules such as DNA, resulting in information on the presence of DNA–protein complexes, even when they do not induce a detectable change in the signals measured with optical tweezers. Moreover, fluorescence microscopy allows counting the number of proteins bound to the DNA. In addition, dynamic processes like diffusion, translocation, and binding kinetics of proteins can be directly monitored using fluorescence imaging. F€ orster resonance energy transfer (FRET) allows distance measurements on the nanometer scale and can be used to study the conformational dynamics of DNA–protein complexes. Simultaneous application of optical tweezers and fluorescence microscopy reaches further than the single-molecule field of biophysics. In recent years, optical tweezers have been applied more and more in vivo. Several research groups have used optical tweezers to hold cells at a fixed position, which was a major breakthrough in single-cell studies, allowing spatial fixing without surface interactions in a sealed device and controlled environment, while enabling continuous observation using advanced imaging techniques (Eriksson et al., 2006; Pang, Song, Kim, Ximiao, & Cheng, 2014). The application of combined optical tweezers and fluorescence microscopy has also been extended to investigate cell-scaffold adhesion (Podlipec & Strancar, 2015), immunological studies and phagocytosis (Tam et al., 2010), permeability as well as molecular partitioning associated with phase transitions in vesicles and lipid bilayers (Bendix & Oddershede, 2011; Bendix, Reihani, & Oddershede, 2010; Kyrsting, Bendix, Stamou, & Oddershede, 2011), the influence of membrane proteins (Brouwer et al., 2015; Prevost et al., 2015), and membrane–cytoskeleton interactions (Leijnse, Oddershede, & Bendix, 2015). In this chapter, we will focus on the broad collection of in vitro singlemolecule assays that have been demonstrated in recent years. We will also discuss several examples of in vivo research providing an impression of the advances in this field as well, albeit without going into detail. We will start with providing a description of the fluorescence and microfluidics techniques that have been combined successfully with optical tweezers. We will put emphasis on the advantages and disadvantages of these combinations, which could serve as guidelines for choosing the most appropriate method

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when designing an experiment. We will also provide an overview of commercial instruments. Next, a collection of recent studies that have combined optical tweezers with fluorescence microscopy will be presented, categorized according to the optical trapping configuration and the fluorescence microscopy technique used. We will discuss examples drawn from across the single-molecule literature, ranging from DNA enzymes, microtubule–protein interactions, DNA/protein conformational dynamics, DNA mechanics, DNA intercalators, lipid–membrane fusion, bacterial motility, and ultimately several FRET studies on nucleosomes and DNA helicases. We also provide a brief protocol of experiments we typically perform in our lab to investigate the binding kinetics of fluorescently labeled proteins or other compounds to DNA, using a combination of dual-trap optical tweezers and wide-field fluorescence microscopy. In this chapter, we aim to highlight the excellent opportunities that the combination of optical tweezers and fluorescence microscopy provides for studying the mechanical properties of DNA (length, flexibility, and elasticity), the kinetics and mechanochemistry of motor proteins, and properties of DNA intercalators.

2. INSTRUMENTATION The combination of optical tweezers and fluorescence microscopy provides a versatile platform that opens up a range of opportunities to gain insight into complex biomolecular transactions. Optical tweezers can hold and move microscopic dielectric particles thanks to transfer of momentum from photons to the object. When forces act on the object, it is pulled out of the laser focus, which results in a deflection of the laser beam that can be detected by a position-sensitive detector. For imaging purposes, a single optical trap can be used to hold objects (Fig. 1A and B). However, in order to visualize and record the tension on a biomolecule, a second anchor is required, which can be provided by a surface or by a second optical trap. In an optical trapping system, it is relatively simple to implement a second optical trapping beam by splitting the trapping light based on polarization (Fig. 1C–F). In the last two decades, optical trapping has been widely employed in different configurations and excellent reviews have been written on the technical aspects of its implementation (Gross, Farge, Peterman, & Wuite, 2010; Moffitt, Chemla, Smith, & Bustamante, 2008; Neuman & Block, 2004). The simultaneous application of optical tweezers and fluorescence microscopy poses several additional technical challenges. For example, the

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Fig. 1 Concurrent optical trapping and fluorescence microscopy. (A) Combination of TIRF fluorescence microscopy with single-trap optical tweezers, where one end of the DNA–protein complex is tethered to an optically trapped microsphere and the other to a surface. Black-thick line: DNA molecule; Black circles: proteins. (B) Combination of confocal fluorescence microscopy with single-trap optical tweezers with one end of the DNA is tethered to an optically trapped microsphere and the other to a surface. (C) Wide-field microscopy combined with dual-trap optical tweezers where both microspheres are being held in two separate optical traps. (D) TIRF microscopy combined with dual-trap optical tweezers with both microspheres being held in two separate traps. The suspended filament (black line) can interact with a single-molecule motor protein (black oval) which is bound to the surface of a pedestal formed on a coverslip. (E) Combination of confocal fluorescence microscopy with dual-trap optical tweezers, where both microspheres are being held in two separate optical traps. (F) Combination of STED fluorescence microscopy with dual-trap optical tweezers with both microspheres being held in two separate optical traps.

light intensities used for the two techniques differ orders of magnitude, which requires efficient separation of trapping and fluorescence light. Moreover, the combination of optical tweezers and fluorescence microscopy is further complicated by optical trap-induced photobleaching of fluorophores (Eggeling, Widengren, Rigler, & Seidel, 1998; Heller et al., 2014; van Dijk, Kapitein, van Mameren, Schmidt, & Peterman, 2004). Despite these challenges a variety of fluorescence microscopy techniques have been successfully combined with optical tweezers (Fig. 1A–F). We categorize the most recent articles combining optical tweezers and fluorescence microscopy in Table 1, according to the optical trap configuration and the fluorescence microscopy technique employed. In this section, we will discuss the strengths and technical difficulties of combining specific fluorescence microscopy techniques with an optical tweezers system. Epi-illuminated wide-field fluorescence microscopy is the simplest fluorescence microscopy technique that has been combined with optical

Table 1 Summary of the Most Recent Literature That Describes Various Combinations of Optical Tweezers and Fluorescence Microscopy Single Trap 1 Trap (Hold)

Wide field

• • •

Eriksson et al. (2006) Pang et al. (2014) Rasmussen, Oddershede, and Siegumfeldt (2008)

1 Trap + Flow

•

Bianco et al. (2001) ✓ Bianco, Bradfield, Castanza, and Donnelly (2007) • Galletto, Amitani, Baskin, and Kowalczykowski (2006) ✓ Handa, Bianco, Baskin, and Kowalczykowski (2005) ✓ Hilario, Amitani, Baskin, and Kowalczykowski (2009) • Pezza, Camerini-otero, and Bianco (2010) • van Mameren, Gross, et al. (2009); van Mameren, Modesti, et al. (2009)

Dual Trap 1 Trap + Surface

• •

Akiyoshi et al. (2010) Ali, Homma, Iwane, Adachi, and Itoh (2004) • Dong, Castro, Boyce, Lang, and Lindquist (2010) • Ferrer et al. (2008) • Fujita et al. (2012) • Funatsu et al. (1997) • Iwaki, Tanaka, Iwane, Katayama, and Ikebe (2006) • Iwaki, Hikikoshi, Ikebe, and Yanagida (2008) ✓ Kudalkar et al. (2015) ✓ Lee, Balci, Jia, Lohman, and Ha (2013) • Lee et al. (2014) • Ngo et al. (2016) • Sarangapani et al. (2014) • Tarsa et al. (2007) ✓ Umbreit et al. (2014)

2 Traps

✓ Arai et al. (1999) ✓ Bao, Lee, and Quake (2003) • Biebricher et al. (2013) ✓ Biebricher et al. (2015) ✓ Brouwer et al. (2015) ✓ Candelli et al. (2014) ✓ Farge et al. (2012) • Farge et al. (2014) • Gross et al. (2011) ✓ King, Gross, Bockelmann, Modesti, and Wuite (2013) ✓ King, Peterman, and Wuite (2016) • Landry, Mccall, Qi, and Chemla (2009) ✓ Mears, Koirala, Rao, Golding, and Chemla (2014) ✓ Min et al. (2009) ✓ Murade, Subramaniam, Otto, and Bennink (2009) • Saito et al. (1994) • van den Broek et al. (2010) ✓ van Mameren et al. (2006) ✓ van Mameren, Gross, et al. (2009) ✓ van Mameren, Modesti, et al. (2009)

2 Traps + Surface

•

Harada, Funatsu, Murakami, Nonoyama, and Ishihama (1999) ✓ Ishijima et al. (1998) • Iwane, Tanaka, Morimoto, Ishijima, and Yanagida (2005) ✓ Komori, Nishikawa, Ariga, Iwane, and Yanagida (2009) ✓ Tanaka, Ishijima, Honda, Saito, and Yanagida (1998) • Tanaka, Homma, White, Yanagida, and Ikebe (2008) • Watanabe et al. (2004)

Miscellaneous

• • • • •

Eriksson et al. (2010) Inman et al. (2014) Mirsaidov et al. (2008) Sasuga et al. (2006) Yogo et al. (2012)

Single Trap 1 Trap (Hold)

Confocal

• • • • • •

STED

Bendix et al. (2010) Bendix and Oddershede (2011) Kyrsting et al. (2011) Leijnse, Oddershede, and Bendix (2015) Podlipec and Strancar (2015) Tam et al. (2010)

1 Trap + Flow

Dual Trap 1 Trap + Surface

✓ Brenner, Zhou, Conway, Lanzano, and Gratton, (2016) • Bornschl€ogl et al. (2013) • Grashoff et al. (2010) ✓ Hohng et al. (2007) • Maffeo, Ngo, Ha, and Aksimentiev (2014) ✓ Ngo, Zhang, Zhou, Yodh, and Ha (2015) • Zhou et al. (2011)

2 Traps

2 Traps + Surface

Miscellaneous

✓ Comstock et al. (2015) • Duesterberg et al. (2015) ✓ Suksombat, Khafizov, Kozlov, Lohman, and Chemla (2015) • Wolfson et al. (2015)

✓ Heller et al. (2013)

The articles in each column of the table are organized according to their optical trap configuration (vertically) and the fluorescence microscopy (horizontally) techniques that are used. The articles within a cell of the table are organized alphabetically based on the last name of the first author. In this chapter, we specifically focus on those combinations that are highlighted with double-line borders. The tick marked articles are discussed in more detail, both the biological assay used as well as a short summary of the pros and cons of the combination of techniques used.

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tweezers (Fig. 1C). It allows for direct visualization of a whole field of view. Moreover, modern electron-multiplying charge-coupled device (EMCCD) or sCMOS cameras with improved signal-to-noise ratios can image single molecules with a temporal resolution in the order of milliseconds (Heller et al., 2014). Illumination of the sample with a relatively wide, parallel light beam often results in high background fluorescence. This limits the concentration of fluorophores in solution at which it is still possible to detect single fluorophores to about 1 nM (Heller et al., 2014). TIRF microscopy significantly reduces the background fluorescent signal. The reflection of the excitation beam on the sample–cover glass interface causes an evanescent field of excitation light that decays exponentially away from the interface, with a typical depth of only a few hundred nanometers inside the sample volume. This substantially reduces background fluorescence, but at the same time requires the experiments to be carried out close to the surface. TIRF is therefore often combined with a trapping configuration where the biomolecule is tethered between a surface and an optically trapped bead or in between two beads that are brought close to a pedestal in the flow chamber (Fig. 1A and D). A disadvantage of the surface-tethered configuration is that the biomolecule is not homogeneously illuminated due to the angle it makes with the interface. Furthermore, measuring close to a surface can sometimes interfere with the biological processes under investigation. Alignment of this fluorescence microscopy technique is only slightly more challenging than an epi-illuminated wide-field fluorescence microscope and commercial implementations in inverted microscopes exist (see later). Confocal fluorescence microscopy has the advantage that background fluorescence can be substantially reduced by spatial filtering of the fluorescence signal using a pinhole, reducing the out-of-focus fluorescence background. In this way, single molecules can be detected in a solution containing about 100 nM fluorophores. This technique, however, requires much more precise alignment and is therefore substantially more challenging to combine with optical tweezers. Visualizing the whole sample requires scanning, since fluorescence is only detected in a very small volume of the sample. Without implementing scanning, confocal fluorescence microscopy has been applied mostly to measure FRET from a single molecule at a fixed position (Fig. 1B and E). Such FRET measurements have a subnanometer spatial resolution (Hohng et al., 2007) and are therefore very suitable to measure conformational changes of biomolecules.

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Scanning confocal fluorescence microscopy (also including STED microscopy) has also been combined with optical tweezers. Scanning the confocal spot over the sample is required to generate an image of part of the sample, which prevents continuous visualization of the whole sample. When detecting single molecules, this scanning approach is typically slower than using camera-based wide-field fluorescence approaches. To overcome this problem, Heller et al. developed a one-dimensional scanning approach, where the two optical traps are being used to linearize the biomolecule (in this case DNA), such that only one-dimensional scanning is necessary (Heller et al., 2013). Implementation of one-dimensional STED allowed for a fivefold enhancement of the diffraction-limited spatial resolution (to
50 nm). This higher spatial resolution allows STED to measure at higher fluorophore concentrations, both in solution and on the DNA itself. A drawback of the 1D-STED approach is the high laser intensity required to generate stimulated emission, substantially increasing photobleaching. Smart and creative application of microfluidic chips that contain micrometer-to-millimeter-sized channels has also led to a wealth of new experimental possibilities. Two main experimental schemes can be distinguished (Fig. 2). In the first approach, buffers are inserted into the microchannels in a sequential manner (Fig. 2A), which allows to construct complex structures such as biomolecules that are attached with one end to a surface and, subsequently, with the other to a bead. In the second approach, buffers are inserted in a flow cell side by side without barriers in between (Fig. 2B). Because the flow is laminar, mixing of buffer flows does not occur. This implementation of microfluidic chips is particularly powerful in optical tweezers configurations without surface tethers, since it allows rapid switching of the buffer composition simply by moving the trap(s) to another laminar “lane.” This rapid switching can, for example, be exploited to suppress high background-fluorescence signals by imaging in a flow channel containing no fluorophores, after incubating in another channel containing the fluorophore. The unique power provided by these sophisticated single-molecule instruments makes studying biological processes almost straightforward. It can, however, be daunting to decide which technique or equipment to use and to know how to deal with technical complexities and maintenance of the instruments. Access to commercial alternatives will undoubtedly make this powerful and versatile combination of techniques accessible to a broad range of researchers in biophysics, biochemistry, drug discovery,

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Fig. 2 Illustration of different geometries of microfluidic chips used in optical tweezers. (A) Sequential delivery of buffers into the microchannel. This configuration allows for the construction of complex structures such as biomolecule attachment to a surface with one end and subsequently attachment to a bead with the other end. Arrows indicate the direction of flow. (B) Insertion of buffers in a parallel fashion. This geometry allows for rapid switching of buffer composition by moving the traps to another laminar “lane.” This configuration enables imaging both in the same lane with labeled entities still present (arrow 1) and in a different lane where no fluorophores are present in the solution (arrow 2).

toxicology, and many other fields. One of the few instruments combining both optical tweezers and fluorescence microscopy techniques and being commercially available is the JPK Nano Tracker™. The optical tweezers system from JPK instruments is designed around a standard inverted microscope and can be combined with various fluorescent techniques. More recently, LUMICKS introduced the C-Trap™. This instrument combines optical tweezers, confocal fluorescence microscopy or STED nanoscopy, and an advanced microfluidics system in a fully integrated configuration. With the development of commercial fluorescence techniques, it is to be expected that single-molecule methods will find wider application in the life and biomedical sciences, providing access to a novel way of studying biomolecular processes in a quantitative way.

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3. APPLICATIONS In the following we will discuss applications of different combinations of optical tweezers and fluorescence microscopy and highlight their capabilities and key practical aspects.

3.1 Single Optical Trap One of the main optical trapping geometries used to study biomolecular interactions is single-trap optical tweezers. Over the past two decades, several studies have employed different configurations of fluorescence microscopy combined with a single-trap geometry, which has yielded valuable knowledge about biomolecular transactions (Table 1). Here, we will first describe some of these studies in which a single trap was used in combination with flow stretching and wide-field microscopy. Next, we will focus on studies in which one end of the biomolecule was tethered to an optically trapped microsphere and the other to a surface, while being visualized using wide-field and TIRF microscopy, a geometry that provides the simplest possible layout that facilitates force and displacement measurements. Finally, we will highlight how confocal microscopy in combination with the single-trap/surface geometry can advance our knowledge of DNA/protein conformational dynamics by enabling FRET measurements. 3.1.1 DNA Enzymes A tremendous amount of new insights in DNA repair enzymes has been obtained using a DNA attached from one side to an optically trapped bead and stretched by buffer flow (see Table 1). Handa et al. described a procedure to track directly DNA translocation by the Escherichia coli RecBCD helicase enzyme (Handa et al., 2005). To visualize a rapidly moving RecBCD molecule directly, a 40-nm streptavidin-coated fluorescent bead (nanoparticle) was attached to RecBCD that had been biotinylated in vivo at a unique site on RecD. The biotinylated DNA was separately bound to a streptavidin-coated polystyrene bead and then mixed with the RecBCD that had been tagged with the fluorescent bead. The bead–DNA–RecBCD– nanoparticle complex was captured by an optical trap and then moved to a second channel of a flow cell containing ATP to start translocation. In their optical trap a Nd:YLF infrared laser (wavelength 1047 nm, 500 mW; Spectra-Physics) was used, which was focused through an oil-immersion

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objective lens (Plan Fluor 100 , 1.3 N.A.; Nikon), to a position 10  15 μm below the upper surface of the flow cell. The trapping was performed in a 4000 μm wide flow cell. The fluorescent nanoparticle– RecBCD complex was excited with an appropriate filter set (Ethidium Bromide set 41006; Chroma Technology Corp.) and the images were captured in real time by an electron bombardment CCD camera (EB-CCD C719023; Hamamatsu Photonics, Hamamatsu, Japan). The position of the nanoparticle relative to the microsphere in their experiments was determined from individual video frames and velocities were determined. This assay allowed determination of the fast-moving rate of RecBCD (up to 1835 bp/s; 0.6 μm/s). Furthermore, the translocation rate of RecBCD was determined on a DNA containing two χ-loci, a recombination hot spot recognized by RecBCD, which was known to reduce the translocation speed of RecBCD. Using this substrate, RecBCD pauses were observed at specific locations, indicating that RecD did not dissociate but underwent conformational modification upon interacting with a χ-locus. This approach can be generally applied to other rapidly translocating motor proteins on DNA. Using a slightly different approach, Bianco et al. (2007) visualized in real time the DNA network formation and cross-bridging activity of RAD54 oligomers, a DNA repair and recombination enzyme in eukaryotes. This was achieved by manipulating two fluorescently labeled (fluorescent dye YOYO-1) DNA–bead complexes trapped side by side, using two optical tweezers in a flow cell. Preformed RAD51–RAD54 nucleoprotein filaments and ATP were introduced by moving the two DNA tethers to a second channel, initiating pairing of the two DNA molecules. These measurements revealed that RAD54 oligomers possess a unique ability to cross bridge or bind double-stranded DNA molecules positioned in close proximity, stimulating the formation of DNA networks, and priming the DNA for rapid and efficient DNA-strand exchange by the recombinase RAD51. In their experiment, optical traps were formed by passing an Nd:YVO4 infrared laser (wavelength 1064 nm, 5 W, Spectra-Physics) through a polarizing beam splitter, which results in fixed and mobile optical traps in the focal plane of the microscope. The resulting, independent laser beams were focused through an oil-immersion objective lens (Plan Apo 100 , 1.4 NA, Leica) to a position 20 μm above the lower surface of the flow cell. Fluorescent DNA–bead complexes were excited using an XCite120 LED (XFO) in combination with a GFP-endow fluorescence cube bandpass filter (Chroma Technology, VT). Finally, fluorescence

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images were captured at video rate by an EB-CCD camera (Hamamatsu) and recorded on digital videotape (DV184) using a digital VCR (Sony DSR-11). Hilario et al. (2009) also used a fluorescent RAD51 to visualize nucleation, assembly, and disassembly of individual nucleoprotein filaments directly. Using this approach, they revealed that the rate of RAD51-nucleoprotein growth increases with a third-order dependence on RAD51 concentration. They determined that a minimum of two to three monomers of RAD51 is required to form a stable nucleus, based on the RAD51-concentration dependence of nucleation. 3.1.2 Interactions of Proteins With Microtubules Combinations of optical trapping with fluorescence microscopy have made important contributions to mitosis and meiosis fields (Akiyoshi et al., 2010; Sarangapani et al., 2014), and in particular to how kinetochores—the molecular machines that drive chromosome separation—are attached to microtubules. To this end, a combination of the single-trap/surface geometry and TIRF microscopy was used by Umbreit et al. (2014) to investigate how Dam1, one of the kinetochore subcomplexes, influences kinetochore– microtubule attachment. They quantified the residence time (5.3  0.5 s) and diffusion (0.021  0.001 mm2s1) of Dam1 complex on microtubules using TIRF microscopy. In addition, they determined the strength of microtubule attachments (7.4  0.4 pN) using an optical trap-based rupture force assay, in which they decorated a polystyrene bead with the Dam1 and allowed it to interact with one side of a microtubule that was attached to the surface with the other side. In a similar approach, Kudalkar et al. (2015) aimed to understand force transmission throughout the kinetochore and to discern the precise role of two other kinetochore subcomplexes, MIND and Ndc80. To this end, they used a two-color TIRF setup with a far-red laser (FTEC-635-0-25-PFQ, Blue Sky Research, Milpitas, CA, USA) that excites either Cy5 or Alexa647, and a blue laser (473-30, LaserPath Technologies, Oviedo, FL, USA; or, more recently, Sapphire 488-75, Coherent, Santa Clara, CA, USA) for the excitation of GFP-tagged kinetochore subcomplexes. They found that MIND dramatically reduced the microtubule-binding time of Ndc80 (Fig. 3). In contrast, MIND alone did not interact with microtubules, even when added at high concentrations, indicating that MIND activates the microtubule-binding activity of the Ndc80 complex. By means of a rupture force assay with a constant loading rate, the strength of the MIND–Ndc80

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Fig. 3 Top panel: Effect of MIND subcomplex on microtubule-binding time of Ndc80. Left: Representative TIRF kymographs of Ndc80c-GFP, Ndc80c-GFP plus Dam1c, MIND-GFP/Ndc80c, and MIND-GFP/Ndc80c plus Dam1c. Right: Diagrams denote each GFP-tagged complex (ovals) and untagged (circles) complex binding to microtubules in kymograph on left. Bottom panel: Tolerance of the MIND/Ndc80c linkage for substantial load. (A) Representative traces of bead position versus time for 20 nM MIND-His/ Ndc80c-FLAG beads under 1.7–2.5 pN of force applied in the direction of microtubule assembly. (B) Survival probability as a function of force is shown for Ndc80c-His, MINDHis/Ndc80c-FLAG, and MIND-His beads with Ndc80c-FLAG and Dam1c-FLAG. The average rupture forces derived from the distributions of survival probability versus force are indicated. Ndc80c-His and MIND-His/Ndc80c-FLAG were not significantly different. Adapted with permission from Kudalkar, E. M., Scarborough, E. A., Umbreit, N. T., Zelter, A., Gestaut, D. R., & Riffle, M. (2015). Regulation of outer kinetochore Ndc80 complex-based microtubule attachments by the central kinetochore Mis12/MIND complex. Proceedings of the National Academy of Sciences of the United States of America, 112, E5583–E5589.

linkage was probed. By performing a dual-label TIRF experiment, Kudalkar and coworkers demonstrated that MIND does not enhance Ndc80 oligomerization, but induces a conformational change in the Ndc80 complex, activating the microtubule-binding domains.

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3.1.3 DNA Helicases The configuration of a single-trap optical tweezers and TIRF discussed above has also been used by Lee et al. (2013) to study dynamics of the E. coli helicase/translocase, UvrD, on a long ssDNA substrate. For this experiment, an oil-immersion objective lens (100 , N.A. 1.40, Olympus) was used for objective-type TIRF microscopy and optical trapping. Fluorescence excitation was only provided close to the cover glass, sample interface using a 532-nm diode-pumped solid-state laser (Spectra-Physics). Fluorescence was detected with an EMCCD camera (Andor iXon). Optical trapping light was provided by a 1064-nm Nd: YAG laser (Spectra-Physics). Using this instrument, they demonstrated UvrD binding to and translocation along ssDNA by tracking the position of individual UvrD fluorescent spots and determined the number of UvrD monomers from fluorescence intensity and counting the number of photobleaching steps. In addition, using the force measured from the optical tweezers, they monitored the unwinding activity. They determined that the processivity of UvrD translocation along ssDNA is 1260 (60) nt, with a velocity of 193 (2) nts1. UvrD monomer translocation stopped at an ssDNA/dsDNA junction, indicating that the translocating UvrD monomers do not make a transition to unwinding duplex DNA.

3.2 Dual-Trap Optical Tweezers The use of two optical traps has several advantages over the single optical trap configurations. First, since there is no need for surface tethering, measurements can be performed far from possibly interfering surfaces. Second, a microfluidic device can be used to change buffer conditions rapidly (Beebe, Mensing, & Walker, 2002; Squires, 2005). Third, since both traps can be positioned independently, the user has threedimensional control over the biomolecular construct that is trapped. Fourth, the dual-trap configuration also has advantages for imaging. Since the traps can be positioned horizontally with respect to the imaging plane, homogeneous illumination is more easily obtained. This makes it possible to perform epi-illuminated wide-field, TIRF, (scanning) confocal as well as STED experiments. These different configurations have been adopted widely (Table 1). Here we will focus on several applications of the dual-trap optical tweezers approach in combinations with fluorescence microscopy.

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3.2.1 DNA Mechanics The dual-trap configuration has been broadly adopted to study the mechanics of biomolecules tethered between two optically trapped beads. Arai et al. used this approach to tie knots in actin and DNA in a high viscosity medium that prevented polymers relaxations (Arai et al., 1999). They were able to estimate knot diameters by comparing the fluorescence intensity of a knot with that of the unknotted polymer. In a similar assay, Bao et al. studied knot behavior in DNA (Bao et al., 2003). They were able to tie several types of knots in the DNA and observed and quantified the diffusion of knots along the DNA. Also other aspects of dsDNA mechanics have been studied extensively. In particular, force–extension behavior has been studied using optical tweezers alone (Gross et al., 2011; Moffitt et al., 2008; Neuman & Block, 2004; Smith, Cui, & Bustamante, 1996), revealing that DNA acts as an entropic spring at forces below 2 pN, followed by an enthalpic regime where the DNA is extended. At forces above 65 pN, DNA overstretches and only a small rise in force is required to extend the DNA to 1.7 times its length. The molecular nature of this overstretching transition had been fiercely debated. Combinations of fluorescence microscopy and optical tweezers have helped settling the debate, thanks to the local information provided by fluorescence microscopy (King et al., 2016; Le, Liu, Lim, & Yan, 2016). In particular, to study the local DNA configuration during overstretching, van Mameren et al. used dual-color epi-illuminated wide-field fluorescence microscopy to image intercalating dyes specific for dsDNA, and ssDNA-binding proteins for ssDNA (van Mameren, Gross, et al., 2009). The DNA construct was attached to the beads with the two opposite strands allowing the DNA to rotate freely with respect to the trapped beads. The intercalator-labeled fraction of the DNA, usually forming a single, continuous patch in the DNA, decreased linearly with extension. The remainder of the DNA was labeled by ssDNA-binding protein, indicating that overstretching can be due to force-induced melting of the dsDNA, resulting in unpeeling of the DNA from the DNA ends. By applying a gentle flow perpendicular to the DNA, they could also observe the ssDNA being flow stretched, thereby confirming their hypothesis. In a follow-up study, King et al. performed experiments on torsionally unconstrained dsDNA without free ends, using fluorescent reporters for ssDNA and dsDNA (King et al., 2013). Under high-salt conditions, during overstretching, part of the

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DNA was labeled by neither reporter, indicating that a third form of DNA, S-DNA, is involved. S-DNA is a DNA conformation proposed before, in which base pairing is still intact but the helicity is largely lost. Under low-salt conditions, patches of the DNA (corresponding to regions with low AT content) could be stained by the ssDNA probe, indicating that local melting bubbles formed. These results directly show that the overstretching of dsDNA can have three different products, peeled ssDNA, ssDNA melting bubbles and S-DNA, and that the ratio between these three depends on sequence, the presence or absence of free DNA ends, and salt concentration. More recently, King et al. determined the mechanics of torsionally constrained dsDNA (King et al., 2016). They found evidence for an additional, previously predicted DNA conformation, P-DNA, with the phosphate backbone facing inwards and the bases outwards, and which is substantially overwound. From the fluorescence data, the percentage of DNA in the P-DNA form could be estimated to be 20–30% which is in good agreement with a previous prediction (Leger et al., 1999). 3.2.2 DNA Intercalators DNA intercalators, which bind between adjacent base pairs of dsDNA, are often used as a fluorescent marker for DNA visualization. It is, however, well known that intercalators perturb the structure and mechanical properties of DNA. To elucidate the effect of intercalators on DNA and to test under what conditions they can be used as DNA probes, Murade et al. and Biebricher et al. combined dual-trap optical tweezers and epi-illuminated wide-field fluorescence microscopy (Biebricher et al., 2015; Murade et al., 2009). Both studies showed that there is a linear correlation between DNA extension and total intercalator fluorescence intensity, indicating that intercalator binding increases with tension. Biebricher et al. quantified the dsDNA-binding and -unbinding kinetics of several intercalators (Biebricher et al., 2015). A combination of force and fluorescence measurements allowed determination of intercalator coverage over an unprecedented four-orders-of-magnitude range. They showed that the force dependence of intercalator binding is mostly governed by the strongly force-dependent unbinding rate (Fig. 4). The unbinding rate varies over seven orders of magnitude, depending on intercalator species, salt conditions, and DNA tension. This detailed quantification of intercalator-binding and -unbinding kinetics provides the insights needed for researchers to select the optimal intercalator species and conditions for a given application.

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5 µm

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Fig. 4 Force-dependent DNA intercalation of a mono- (SbG, open symbols) and a bisintercalator (YOYO, solid symbols) at 1000 mM NaCl. (A) DNA elongation and representative fluorescence images as a function of tension. (B) Binding constant as a function of tension, calculated using elongation data. Adapted with permission from Biebricher, A. S., Heller, I., Roijmans, R. F. H., Hoekstra, T. P., Peterman, E. J. G., & Wuite, G. J. L. (2015). The impact of DNA intercalators on DNA and DNA-processing enzymes elucidated through force-dependent binding kinetics. Nature Communications, 6, 1–12.

3.2.3 DNA Enzymes Dual-trap optical tweezers also allow quantitative analysis of DNA-binding proteins. One such protein that has frequently been studied this way is RAD51, which forms ATP-dependent filaments on dsDNA and ssDNA. van Mameren et al. used a combination of optical tweezers and epiilluminated wide-field fluorescence microscopy to study the mechanics and kinetics of RAD51 filaments on dsDNA (van Mameren et al., 2006). They showed that, in the absence of ATP hydrolysis, RAD51 forms immobile filaments that remain stably bound to DNA even at high forces. Moreover, the measurements allowed estimation of the dsDNA length increase upon binding RAD51 to be 150%. Notably, the filaments of RAD51 on DNA are very rigid and could hardly be stretched when the DNA molecule is pulled (Fig. 5).

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Fig. 5 Elastic properties of a single 48 kbp λ-dsDNA molecule partly coated with fluorescent RAD51. (A) Fluorescence image (left) of such an assembly, tethered between two streptavidin-coated polystyrene beads. Kymograph (right) generated from the successive frames of the movie recorded during extension of the construct. The corresponding force time trace is depicted in gray scales (top bar; white corresponds to 90 pN). (B) Force–extension curve corresponding to the construct in (A). The gray trace shows a bare λ-DNA reference curve. (C–E) Force–extension curves of the bare zone (i), the continuous fluorescent zone (ii), and the composite fluorescent zone (iii) as indicated on the right of the kymograph. Adapted with permission from van Mameren, J., Modesti, M., Kanaar, R., Wyman, C., Wuite, G. J. L., & Peterman, E. J. G. (2006). Dissecting elastic heterogeneity along DNA molecules coated partly with Rad51 using concurrent fluorescence microscopy and optical tweezers. Biophysical Journal, 91(8), L78–L80.

Using a similar approach, Candelli et al. focused on RAD51-filament nucleation and growth on both dsDNA and ssDNA (Candelli et al., 2014). First, by counting the number of nuclei that formed in a given time, they determined the nucleation rate, which increased with RAD51 concentration. From the fluorescence intensity of individual nuclei, they determined the number of RAD51 monomers per nucleus and found a wide distribution of nucleus sizes. In addition, filament growth was studied with a single-molecule fluorescence recovery after photobleaching assay, in which nuclei were first completely photobleached, followed by incubation of the DNA in a RAD51-containing buffer. When a new fluorescent spot occurred at the same location as a previously photobleached one, this was interpreted as a growth event. From such experiments the filament growth rate was determined. Remarkably, both nucleation and growth rates are force dependent for RAD51 binding to dsDNA, but not for binding to ssDNA.

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van Mameren et al. looked at the disassembly of RAD51 filaments from dsDNA (van Mameren, Modesti, et al., 2009). Preformed RAD51nucleoprotein filaments were rapidly brought from a nonhydrolyzing to an ATP-hydrolyzing condition by moving them to another channel in the flow cell. While holding the traps at a fixed distance, tension was observed to increase, while the RAD51 fluorescence intensity decreased, consistent with RAD51 disassembling from the DNA (Fig. 6). Remarkably, a sudden release of the tension resulted in an enhanced disassembly rate. This result was interpreted to indicate that the force-dependent event was not ATP hydrolysis, but the actual detachment of a RAD51 monomer from the filament. Dissociation occurred in bursts interspersed

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Fig. 6 RAD51 disassembly rate is reversibly reduced by DNA tension. Top: Kymograph of a RAD51–dsDNA complex, held at fixed length. Bottom: Intensity trace (red, descending line from top left) and tension trace (blue, ascending line from bottom left) of a RAD51–dsDNA complex. Tension-stalled disassembly is reinitiated by tension release (orange dashes). Adapted with permission from van Mameren, J., Gross, P., Farge, G., Hooijman, P., Modesti, M., Falkenberg, M., et al. (2009a). Unraveling the structure of DNA during overstretching by using multicolor, single-molecule fluorescence imaging. PNAS, 106(43), 18231–18236; van Mameren, J., Modesti, M., Kanaar, R., Wyman, C., Peterman, E. J. G., & Wuite, G. J. L. (2009b). Counting RAD51 proteins disassembling from nucleoprotein filaments under tension. Nature, 457(7230), 745–748.

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with pauses comparable to the typical RAD51-catalyzed ATP hydrolysis time, providing support for a model in which RAD51 disassembly only takes place from filaments ends after ATP hydrolysis in the RAD51 monomer on the end. Another DNA-binding protein that has been studied using optical tweezers and wide-field fluorescence microscopy is TFAM, a key protein in mitochondrial DNA transcription and compaction (Farge et al., 2012). Fluorescence intensity measurements of dsDNA saturated with fluorescently labeled TFAM allowed determination of the footprint of the protein. The intensity of individual fluorescent spots observed at low TFAM concentrations indicated that TFAM binds as a monomer to dsDNA. These monomers diffused along the DNA and with single-molecule tracking the diffusion coefficient was determined. Force–extension measurements at different TFAM concentrations showed that TFAM binds to DNA in a cooperative way, decreasing the persistence length. In a follow-up study, it was shown how compaction by TFAM blocks unwinding of duplex DNA resulting in inhibition of mitochondrial DNA transcription and replication (Farge et al., 2014). To obtain even higher-resolution insights in TFAM diffusion and DNA binding, Heller et al. applied optical tweezers in combination with STED super-resolution microscopy (Heller et al., 2013). The advantage of using STED was that TFAM could be studied at higher, more physiologically relevant concentrations in solution and on the DNA. It was observed that TFAM monomers can cluster and form multimers, diffusing with a lower diffusion constant (Fig. 7).

3.2.4 Lipid–Membrane Fusion Using a very different dual-trap optical tweezers assay, Brouwer et al. studied lipid–protein interactions (Brouwer et al., 2015). To this end, they coated the two trapped beads with lipid bilayers and repeatedly brought them together and separated them. In the presence of Doc2b, a protein that is involved in SNARE-mediated membrane fusion during neurotransmittervesicle release, high rupture forces were observed. These high rupture forces indicated Doc2b-mediated membrane fusion. When the membranes were fused, fluorescently labeled lipids on one bead could diffuse to the other bead, while dyes in the lumen between membrane and bead did not, indicating that Doc2b mediates fusion of only outer leaflet of the lipid bilayer, known as hemifusion (Fig. 8).

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Fig. 7 Imaging protein dynamics at high TFAM protein density, using STED highresolution microscopy. (A) Kymographs of TFAM-Atto 647 N dynamics on λ DNA (F ¼ 4 pN). Scale bars, 1 μm. The DNA is in reductive–oxidative system buffer, and at t ¼
66 s, the STED beam is switched on at 6 mW (right box in the kymograph (red) borders; FWHM ¼ 75 nm). Proteins that diffuse within diffraction-limited regions are indicated with arrows. (B) Top panel: a kymograph of a TFAM-Atto 647 N oligomerization event. Bottom panel: the MSD analysis of trajectories i–iii as indicated in top panel (error bars: standard deviation; mean number of lines ¼ 130). Adapted with permission from Heller, I., Sitters, G., Broekmans, O. D., Farge, G., Menges, C., Wende, W., et al. (2013). STED nanoscopy combined with optical tweezers reveals protein dynamics on densely covered DNA. Nature Methods, 10(9), 910–916.

3.2.5 Cytoskeletal Motor Proteins Dual-trap optical tweezers in combination with fluorescence microscopy have also been used to study the interaction of myosin motor proteins with filamentous actin. To this end, myosin motors were attached to a pedestal in the flow cell, while an actin filament was held between two optically trapped beads. Using this assay, interactions and force generation

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Fig. 8 Investigation of lipid–protein interaction using dual-trap optical tweezers combined with epi-illuminated wide-field fluorescence microscopy. (A) In the presence of fluorescent NBD-PE in a single membrane-coated bead, membrane-stalk formation was accompanied by a strong fluorescence increase in the unlabeled membrane and a concurrent decrease in the labeled membrane, indicative of either hemifusion or full membrane fusion. (B) To distinguish between hemifusion and complete fusion, content mixing was tested by coating the beads with nonfluorescent PC/PS liposomes, while one of the beads was loaded with fluorescein in the liposomal lumen. On a timescale of 1000 s, fluorescence increase in the unlabeled bead or membrane stalk was not observed, indicative for hemifusion. (C) Schematic representation of the hemifused configuration with Doc2b bound to the membrane surface. (D) Fluorescence images of bead pair in (A) at three different time points. Scale bar: 1 μm. (E) Fluorescence images of bead pair in (B) at three different time points. Scale bar: 1 μm. Adapted with permission from Brouwer, I., Giniatullina, A., Laurens, N., Weering, J. R. T. Van, Bald, D., Wuite, G. J. L., et al. (2015). Direct quantitative detection of Doc2b-induced hemifusion in optically trapped membranes. Nature Communications, 6, 1–8.

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can be measured with the tweezers, while binding and release of fluorescent ATP analogues to the motor proteins can be followed with TIRF microscopy. With this approach, step sizes, processivity, and ATPase activity of several myosin motors were characterized (Ishijima et al., 1998; Iwane et al., 2005; Komori et al., 2009; Tanaka et al., 1998, 2008; Watanabe et al., 2004). 3.2.6 Bacterial Motility Min et al. and Mears et al. held a living bacterium fixed with two optical traps, while flagellum position was monitored using fluorescence and cellbody displacement with the tweezers (Mears et al., 2014; Min et al., 2009). Fourier analysis of the cell-body displacements revealed two frequencies, one corresponding to rotation of the flagellum and the other with the resulting rotation of the bacterium. The bacterial rotation signal contained indications of periods of constant, directional movement, and tumbling, consistent with two different kinds of appearances of the flagellum fluorescence images (Min et al., 2009). Improvements in fluorescence imaging allowed Mears et al. to distinguish multiple flagella at the same time and determining their sense of rotation (Mears et al., 2014). They demonstrated that only a single flagellum rotating clockwise was required to switch the bacterium into tumbling mode (Fig. 9). They also showed that flagella do not switch their sense of rotation independently (as was assumed before),

Fig. 9 Assessment of bacterial motility. Top: Representative images from a trapped cell with three flagella. The approximate location of the unlabeled cell body is indicated by a dashed line. Flagella rotating CW (purple) and CCW (underlined-white) are numbered in frames in which they appear distinct. Bottom: corresponding cell-body rotation signal for the same cell as detected from deflections of the trapping laser. Tumbles (shaded area) were determined from the erratic cell-body rotation signal. Adapted with permission from Mears, P. J., Koirala, S., Rao, C. V, Golding, I., & Chemla, Y. R. (2014). Escherichia coli swimming is robust against variations in flagellar number. eLIFE, 3, e01916.

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but do so in correlated fashion, which has important implications for understanding bacterial motility and chemotaxis.

3.3 FRET Studies With Confocal Fluorescence Microscopy Force-measuring tools like atomic force microscopy or optical tweezers cannot detect small-scale conformational changes, unless a relatively strong force is applied. This limitation has been overcome by combining single-trap optical tweezers with FRET confocal microscopy, providing access to measurements of conformational dynamics. Hohng et al. used this approach to investigate the dynamics of Holiday Junction structures (HJ) (Hohng et al., 2007). An HJ is a four-stranded DNA structure that is an intermediate state during homologous recombination. A trapping beam of 1064 nm was fixed in the field of view of the microscope, while forces were applied by moving the surface-tethered HJ using a piezoelectric sample stage. The confocal laser focus (532 nm) was scanned to follow the motion of the molecule in response to the moving optical trap. By specifically labeling the HJ at different locations a 2D reaction landscape could be obtained, by probing the HJ dynamics in response to pulling forces in three different directions. In this way, global structural information could be obtained on transient species involved in HJ conformational changes. The application of combined optical tweezers and FRET microscopy has also been extended to investigate DNA–protein interactions. For example, Ngo et al. (2015) employed single-trap optical tweezers and FRET microscopy to manipulate an individual nucleosome under force and simultaneously probe its local conformational changes. They anchored a DNA molecule with a nucleosome-binding site with one end to a glass surface and with the other end to an optically trapped microsphere. The DNA contained FRET dye pairs at various locations. They found that the DNA, which is wrapped around the nucleosome, unwraps asymmetrically from a nucleosome: from one side unwrapping occurs at low forces (3–5 pN), while from the other side, substantially higher forces are required for unwrapping (12–15 pN). These results have important implications for chromosome remodelers, which move or expel nucleosomes from DNA. In their setup, they applied an infrared laser (1064 nm, 800 mW, EXLSR-1064-800-CDRH, Spectra-Physics) to form the optical trap through the back port of a commercial microscope (Olympus) and applied forces on the sample tethers by moving the microscope slide using a piezoelectric stage (Physik Instrumente). The position of the tethered bead with

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respect to the trap was determined using a quadrant photodiode (SPOT/ 9DMI, UDT). Excitation light for confocal excitation (wavelength: 532 nm, World Star Tech) was coupled into the side port of the microscope and was scanned by a piezo-controlled steering mirror (S-334K.2SL, Physik Instrumente). Fluorescence was filtered using a bandpass filter (HQ580/60m, Chroma) and separated from the excitation light using a dichroic mirror (HQ680/60m, Chroma) before detection by two avalanche photodiodes (APDs). In a different approach, Suksombat et al. studied the interaction of ssDNA-binding protein, SSB, with a short stretch (70 nucleotides) of ssDNA held between two optical traps with dsDNA handles (Suksombat et al., 2015). In their experiments, they used two high-resolution optical traps, formed by timesharing a single IR laser (a 5-W, 1064-nm diodepumped solid-state laser, YLR-5-1064-LP; IPG Photonics), by intermittently deflecting the laser in two directions with an acousto-optic modulator (IntraAction). Constant-force experiments were performed with a PID controller loop that monitored the trapped bead positions and controlled the trap separation to maintain a constant tension on a tethered DNA molecule. Fluorescence probes were excited by a 532-nm 5-mW laser (DPGL-05S, World Star Tech) interlaced with the trapping IR laser at a rate of 66 kHz and imaged onto two APDs (Perkin Elmer). Using a force clamp they observed step-wise length switches of the DNA tether, due to partial wrapping and unwrapping of the DNA around SSB (Fig. 10). This switching could also be observed using FRET between dyes on the DNA and SSB. They also observed changes in FRET efficiency uncorrelated to changes in DNA extension, indicative of SSB diffusing along the DNA. With a similar assay, Comstock et al. studied UvrD helicase unwinding a DNA hairpin (Comstock et al., 2015). Changes in DNA extension revealed unwinding and rezipping of the DNA. Concomitant fluorescence measurements allowed discrimination of activity of UvrD monomers and dimers. For monomers, frequent, repetitive switching between unwinding (of at most 20 bp) and rezipping was observed, while dimers processively unwound the DNA over lengths of 70 bp. FRET experiments showed that the conformation of the protein, open or closed, correlated to unwinding or rezipping activity. Furthermore, the subnanometer sensitivity of FRET in measuring distances, together with piconewton force sensitivity of optical tweezers enables studies of mechanotransduction. With the help of this capability, Brenner et al. used this combination to determine how spider-silk

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Fig. 10 SSB-binding modes and diffusion mechanism. Top: schematic of wrapping, releasing and sliding of fluorescently labeled SSB (SSBf) on ssDNA. Bottom: representative traces showing combined fluorescence and DNA extension measurements. Change in extension (top) and fluorescence (middle) of donor (SSBf) and acceptor (Cy5) are measured simultaneously. Together, FRET efficiency (bottom) and extension change (top) reveal the SSB wrapping states (i and ii, iii and iv) and their dynamics (ssDNA wrapping/releasing and sliding). Adapted with permission from Suksombat, S., Khafizov, R., Kozlov, A. G., Lohman, T. M., & Chemla, Y. R. (2015). Structural dynamics of E. coli single stranded DNA binding protein reveal DNA wrapping and unwrapping pathways. eLIFE, 4, e08193.

flagelliform repeat peptides react to force (Brenner et al., 2016). They first showed that, at zero force, each peptide has a conformation stable on the timescale of one second to minutes. Next, they showed, by measuring FRET efficiency as a function of force, that the peptides behave as linear springs. This was unexpected since disordered proteins act as nonlinear springs. The results indicate that these peptides are highly compact and in an ordered, rod-like coil structure. Brenner et al. also used these peptides as intracellular force sensors, by integrating them in the focal-adhesion protein vinculin and measuring FRET using fluorescence lifetime imaging.

4. EXPERIMENTAL PROTOCOL In this section, we describe a general scheme to perform experiments such as those described in the previous section using a dual-trap optical tweezers setup combined with wide-field epi-fluorescence microscopy, as we typically perform them in our laboratory (Biebricher et al., 2015; Farge et al., 2012; King et al., 2016).

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4.1 Experimental Setup A detailed description of our dual-trap optical tweezers setup combined with epi-fluorescence microscopy has been presented elsewhere (Gross et al., 2010). In brief, two perpendicularly polarized trapping beams are generated by splitting a 1064-nm trapping laser (YLR-LP, IPG photonics) with a halfwave plate and a polarizing beam splitter. These beams can be manipulated independently using steerable mirrors. Displacement of the beads with respect to the trap centers are measured by back-focal-plane interferometry using two position-sensitive detectors (DL100-7-PCBA3, Pacific Silicon Sensor); the position of the beads is determined using bright-field imaging, using blue LED illumination and a CCD camera (both Thorlabs). Fluorescence excitation is provided, for example, by a 532-nm diode-pumped solid-state laser (Samba, Cobolt) into the microscope and the fluorescence is detected using an EMCCD camera (Ixon3 897, Andor). A microfluidic chip containing up to six channels, forming parallel flow lanes (Fig. 2B), is mounted on the microscope stage and connected to a flow system based on pressurized air (u-Flux, LUMICKS). Valves are used to control independently the buffer flow in separate channels. The DNA construct is obtained by binding of biotinylated nucleotides to the 30 -recessive ends of Bacteriophage Lambda (λ) DNA (
48.5 kb) (Roche) according to protocols described before (Gross et al., 2010).

4.2 Methods •

•

•

•

Cleaning of the microfluidic flow cell: rinse the flow cell thoroughly with bleach (a sodium hypochlorite solution) to clean the surface. Rinse afterward with sodium thiosulfate solution to neutralize remaining bleach and subsequently with water. Passivation of flow cell to prevent sticking of proteins to surface: flush the flow cell with casein or BSA solution, followed by a Pluronic© F127 solution. Introduction of buffer solutions: flush a dilution of streptavidin-coated 4.5-μm microspheres (Spherotech) through the first microfluidic channel. Flush a buffered solution of the biotinylated DNA construct through the second channel and buffer through the third channel. Flush your fluorescently labeled protein or compound of interest through the fourth and/or the fifth channel. Catching DNA: trap, in the presence of flow, two beads by moving the traps to the first channel. Subsequently, move the traps to the second

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•

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flow channel and monitor jumps in force that indicate binding of DNA to one of the beads. Move the beads to the third channel and approach and withdraw the downstream bead from the upstream bead until a change in force indicates the formation of a DNA dumbbell construct. Protein incubation: move DNA construct to the channel that contains your fluorescently labeled protein or compound of interest and allow it to bind to the DNA. Depending on the binding kinetics, this will take seconds up to several minutes. Note that for some compounds, binding is facilitated by application of a pulling force on the DNA. Single-molecule fluorescence: move the incubated DNA back to the third channel containing only buffer to reduce the background fluorescence. Illuminate with fluorescence excitation laser and measure fluorescence images with the EMCCD camera. In this way the unbinding kinetics of the compound or protein of interest can be followed, for example, as a function of force.

5. CONCLUSION The state-of-the-art combination of optical tweezers with fluorescence microscopy and microfluidics techniques provides a valuable addition to the single-molecule toolkit. It has enabled scientists to study biological systems with previously unobtainable precision and clarity on a vast number of different biological interactions as illustrated in this chapter. Some of the most recent advancements such as STED microscopy promise to give rise to even more exciting new possibilities by pushing the experimental limits toward single-molecule experiments in conditions closer to the physiological reality. Moreover, the current state of maturity of the technology and the concurrent development of their application in biological assays, as is reviewed in this chapter, paves the road for broadening the application of this combined technique from the realm of fundamental research in a specialized biophysics laboratory to biochemical, biological, and pharmaceutical laboratories. Finally, commercial solutions are becoming increasingly available, which makes this powerful combined approach accessible to a broad range of researchers from different backgrounds. When this technology spreads we expect to see more innovative and exciting single-molecule experiments which can revolutionize research in biophysics, biochemistry, drug discovery, toxicology, and many other fields.

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ACKNOWLEDGMENTS This work was supported by an STW-HTSM grant (to W.H.R. and G.J.L.W.), VICI grants (to G.J.L.W. and E.J.G.P.), a VIDI grant (to W.H.R.) of the Nederlandse Organisatie voor Wetenschappelijk Onderzoek (NWO), and a FET Open grant from Horizon 2020 program of the European Union. Disclosure: G.W. and E.P. are cofounders and have financial interested in LUMICKS B.V.

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CHAPTER FIVE

Direct Visualization of Helicase Dynamics Using Fluorescence Localization and Optical Trapping C.-T. Lin*, T. Ha*,†,1 *Johns Hopkins University, Baltimore, MD, United States † Howard Hughes Medical Institute, Baltimore, MD, United States 1 Corresponding author: e-mail address: [email protected]

Contents 1. Introduction 2. Materials 2.1 DNA 2.2 Reagents and Buffers 2.3 Single-Molecule Fluorescence-Force Spectroscopy 2.4 Data Acquisition and Analysis 3. Methods 3.1 PEG-Passivated Slide/Coverslip Preparation 3.2 Sample Chamber Assembly 3.3 Single-Molecule TIRF-Optical Tweezers Acknowledgments References

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Abstract Helicases control the accessibility of single-stranded (ss) nucleic acid (NA) generated as a transient intermediate during almost every step in cells related to nucleic acid metabolisms. For subsequent processing, however, helicases need to adjust the pace of unwinding adequately to avoid ssNA exposure to nucleases. Therefore, understanding how the unwinding process of helicases is regulated is crucial to address genome integrity and repair mechanisms. Using single-molecule fluorescence-force spectroscopy with fluorescence localization, we recently observed the stoichiometry of UvrD helicase, which determines the functions of UvrD: translocation and unwinding. For the first time, we provide direct evidence that a UvrD dimer is required to initiate the unwinding pathway. Moreover, with subpixel precision of fluorescence localization, the dynamic parameters of helicases can be obtained directly. Here, we present detailed single-molecule assays for observing the biochemical activities of helicases in real time and revealing how mechanical forces are involved in protein–nucleic acid interactions. These singlemolecule approaches are generally applicable to many other protein–nucleic acid systems. Methods in Enzymology, Volume 582 ISSN 0076-6879 http://dx.doi.org/10.1016/bs.mie.2016.08.004

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2017 Elsevier Inc. All rights reserved.

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1. INTRODUCTION In the past years, many single-molecule detection techniques were developed which enable us to observe and distinguish directly subpopulations in the dynamics of nucleic acids and proteins and their interactions in real time (Joo, Balci, Ishitsuka, Buranachai, & Ha, 2008; Lu, Xun, & Xie, 1998; Robison & Finkelstein, 2014). In particular, single-molecule fluorescence and optical tweezers are the two most widely used approaches in studying biological molecules (Ha et al., 1996; Moffitt, Chemla, Izhaky, & Bustamante, 2006; Roy, Hohng, & Ha, 2008; Visscher, Gross, & Block, 1996). In contrast to conventional ensemble approaches, single-molecule methods allow us to probe interactions even under nonequilibrium conditions and, therefore, provide a window to explore the heterogeneity among molecules, revealing transitions and different modes of molecular dynamics which are otherwise hidden in ensemble measurements (Greenleaf, Woodside, & Block, 2007; Joo et al., 2008; Walter, Huang, Manzo, & Sobhy, 2008). Helicases are ATP-powered motor proteins and are essential for living cells. Single-stranded, double-stranded, or hybrid nucleic acids are substrates for helicases; besides translocation and unwinding, helicases can remodel their substrates with or without other bound proteins (Jankowsky, Gross, Shuman, & Pyle, 2001; Matson, Bean, & George, 1994; Schmid & Linder, 1992). In other words, helicases are involved in almost every step of nucleic acid metabolism processes (Lohman & Bjornson, 1996; Tuteja & Tuteja, 2004). Two major activities of helicases are translocation and unwinding, which have drawn much attention in the past decade, and studies have focused on the biochemical, structural, and genetic bases for these activities (Donmez & Patel, 2006; Jankowsky & Fairman, 2007; Mackintosh & Raney, 2006; Martin, Mark, & Dale, 2007; Matson et al., 1994; Vindigni, 2007). At the same time, the maturation of single-molecule methodology has provided enough capability to reveal individual helicase activities. However, measuring only either fluorescence or force to characterize helicase activities are not enough to elucidate helicase functions fully. Therefore, we have applied a hybrid platform of single-molecule fluorescence localization under total internal reflection fluorescence (TIRF) configuration combined with optical tweezers to reveal helicase translocation and unwinding in real time. This approach allowed us to image single helicase translocation on ssDNA directly, show that a UvrD monomer is stalled

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at ss/dsDNA junctions until the arrival of a second monomer activates unwinding (Lee, Balci, Jia, Lohman, & Ha, 2013), and image a new mode of translocation for NS3 helicase (Lin et al., 2014). In this chapter, we describe detailed protocols for the experimental assays mentioned earlier, including the materials and steps for sample preparation, and data acquisition and analysis (Lee, 2013). We recommend the readers refer to comprehensive guides elsewhere for assembling a home-built fluorescence-force spectroscopy setup (Lin & Ha, in press).

2. MATERIALS 2.1 DNA 1. ssDNA template for rolling circle amplification (RCA): 50 - AGG AGA AAA AGA AAA AAA GAA AAG AAG G -30 (Lee et al., 2014). 2. Biotinylated primer for RCA: 50 -/biotin/TCT CCT CCT TCT -30 (Lee et al., 2014). 3. 4957-bp double-stranded (ds) DNA PCR-amplified from bacteriophage λ DNA (see Section 3.3.3).

2.2 Reagents and Buffers 1. 2. 3. 4. 5. 6. 7. 8. 9. 10. 11. 12. 13. 14. 15. 16.

T4 DNA ligase (NEB). 25 μM dTTP/dCTP. 20 mg/mL BSA (NEB). Phi29 DNA polymerase (NEB). Digoxigenin-11-ddUTP (Roche). Antidigoxigenin-coated beads (see Section 3.2). Bacteriophage λ DNA (NEB). Terminal transferase (NEB). T7 exonuclease (NEB). Protein G-coated polystyrene beads (Spherotech, 880 nm in diameter). MES (2-(N-morpholino) ethanesulfonic acid, Sigma-Aldrich). EDC (N-(3-Dimethylaminopropyl)-N0 -ethylcarbodiimide) hydrochloride (Sigma-Aldrich). NHS (N-Hydroxysuccinimide, Sigma-Aldrich). Antidigoxigenin (Roche). MES buffer (100 mM MES. Adjust pH to 6.5 with 5 M NaOH). Reconstitution buffer (0.019 M NaH2PO4, 0.081 M Na2HPO4, 0.14 M NaCl, 2.7 mM KCl).

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17. Bead storage buffer (0.039 M NaH2PO4, 0.061 M Na2HPO4, 0.14 M NaCl, 2.7 mM KCl, 0.1 mg/mL BSA, 0.1% (v/v) Tween-20, 0.02% (w/v) sodium azide). 18. Termination buffer (1 M Tris–HCl, pH 6.8). 19. Slide staining jar (with cover, Wheaton). 20. Coverslips (24  40 mm, white borosilicate, VWR). 21. Glass slides (300  100 , 1 mm thick, Erie Scientific). 22. Amino silane (N-(2-Aminoethyl)-3-Aminopropyltrimethoxysilane, United Chemical). 23. 10% alconox. 24. Acetone (99.7%, VWR). 25. 1 M/3 M/5 M KOH. 26. Methanol (99.7%, VWR). 27. Biotin-PEG-Succinimidyl Valerate (Laysan Bio). 28. mPEG-Succinimidyl Valerate (Laysan Bio). 29. Acetic acid (99.7%, Fisher Chemical). 30. 0.1 M sodium bicarbonate buffer. 31. PEG-passivated slides (coverslip and glass slide). 32. Double-sided tape (3 M). 33. Epoxy (All-spec). 34. T50 buffer (10 mM Tris–HCl, 50 mM NaCl, pH 8.0). 35. 0.2 mg/mL neutravidin (Thermo Fisher Scientific, USA). 36. Large orifice tip (VWR). 37. Imaging buffer: 10 mM Tris–HCl, pH 8.0, 0.1 mg/mL BSA, 0.8% (w/v) D-glucose, 165 U/mL glucose oxidase, 2170 U/mL catalase, 3 mM Trolox (-6-Hydroxy-2,5,7,8-tetramethylchromane-2-carboxylic acid), 5 mM MgCl2, 1 mM DTT, 2% (v/v) glycerol and indicated amounts of NaCl, ATP, and helicase.

2.3 Single-Molecule Fluorescence-Force Spectroscopy An extensive description of the assembly and calibration of a home-built hybrid instrument combining TIRF microscopy and optical tweezers can be found elsewhere (Lin & Ha, in press).

2.4 Data Acquisition and Analysis The hybrid setup of TIRF-optical tweezers is controlled by an in-house program written in Microsoft Visual C++. The data is analyzed by MathWorks MATLAB code.

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3. METHODS 3.1 PEG-Passivated Slide/Coverslip Preparation To reduce nonspecific surface adsorption of proteins and other biological molecules in single-molecule experiments, a layer of polymer, polyethyleneglycol (PEG), is used to coat the glass slide and coverslip. The following is the general protocol for preparing PEG-coated slides and coverslips (Selvin & Ha, 2008). 3.1.1 Precleaning 1. Drill a pair of inlet/outlet holes per sample channel on glass slide, and the numbers of sample channel per slide will depend on one’s application (generally four sample channels are recommended for optical tweezers experiments). 2. Scrub the slides with 10% alconox and rinse with distilled water. Make sure that there is no visible residue on the slides. 3. Microwave slides in water for 10 min. 4. Put the slides and new coverslips in separate slide staining jars. 5. Pour acetone in the containers and sonicate for 30 min. 6. Rinse the containers with the slides and coverslips with water. 7. Burn the slides and coverslips with a propane torch and put them back into dry containers. 3.1.2 Aminosilanization 1. Pour 3 M KOH in the slide and coverslip containers and sonicate for 20 min. 2. Rinse with water three times. 3. Rinse with MeOH twice and sonicate the containers with MeOH for 5 min. 4. Pour 150 mL MeOH, 7.5 mL acetic acid, and 1.5 mL aminosilane into a flask. Quickly mix well. 5. Remove MeOH from the containers and pour the solution from the flask into the containers. 6. Incubate 10 min on bench top. Sonicate for 1 min, and incubate another 10 min. 7. Remove the solution in the containers and rinse with clean MeOH at least twice or until there is no acetic acid smell.

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8. Air-dry coverslips, glass slides, and the containers with nitrogen and cover the lids of the containers. 3.1.3 PEGylation 1. Measure 84 mg sodium bicarbonate and dissolve it into 10 mL of water. 2. Dissolve 2 mg biotin-PEG and 100 mg mPEG into 400 μL of sodium bicarbonate buffer. The total amount of PEG solution can be scaled up while the ratio is maintained, 80 μL of PEG solution is required per slide/coverslip set. 3. Mix the solution gently by pipetting and then centrifuge 2 min at 10,000  g. 4. Prepare moisture boxes in which slides can be incubated. Place slides in the box, put 80 μL of the PEG solution on top of the slide, and then sandwich the solution with a coverslip. Moisture box can be either home-made or purchased from Thermo Fisher Scientific (slide moisture chamber). 5. Incubate for at least 5 h and no longer than 12 h to avoid drying out PEG solution. 6. After incubation with PEG solution, clean the PEGylated coverslips and slides with water and air-dry them. Store sets of sample chambers individually in 50-mL tubes with a hole at the end; a single set of sample chamber includes one coverslip and one slide. 7. Vacuum seal tubes in food saver bags and store them at 20°C in a freezer.

3.2 Sample Chamber Assembly 1. Take one 50-mL tube containing a sample chamber set from the 20°C freezer and keep it in the dark until it warms up to RT. 2. Attach two pieces of double-sided tape to the PEG-passivated glass slide in such a way that a 8 mm gap is formed between the two pieces of tape perpendicular to the slide’s long axis. 3. Put a coverslip over the double-sided tape to form a 20 μL volume sample channel. Make sure the PEG-passivated side of the coverslip is facing inside the channel. 4. Seal the gaps, which are parallel to the slide’s long axis, between coverslip and glass slide with epoxy and let sit for 5 min. Leave drilled holes (inlet and outlet) open on glass slide for further buffer flow.

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3.3 Single-Molecule TIRF-Optical Tweezers The TIRF-based measurements are useful for characterizing helicases’ biochemical activities and dynamic motions at zero force. With tension applied to the helicase-bound nucleic acid, one can study the process of helicase unwinding, translocation with its cofactor, and observe how mechanical force regulates the activity of the complexes on nucleic acids. This method has successfully combined fluorescence localization with TIRF-based measurement; therefore, the precision of a single fluorophore position can be as good as 15 nm with 20 ms time resolution (Lee et al., 2013). After aligning the optical trap and the TIR excitation laser beam and following every necessary calibration, we can monitor the movement of fluorescently labeled molecules under desirable mechanical manipulations. A thorough guide of assembling a TIRF-optical tweezers microscope can be found elsewhere (Lin & Ha, in press). 3.3.1 Preparation of the Antidigoxigenin-Coated Beads A detailed protocol to cross-link antidigoxigenin chemically to protein G beads can be found elsewhere (Zhou, Schlierf, & Ha, 2010). Items 10–18 listed under Section 2.2 are required for the reaction. 3.3.2 General Sample Preparation and Data Acquisition Protocol For making the nucleic acid templates for fluorescence-force measurements, we first incubate the premade biotinylated nucleic acid structure (biotin is added to the nucleic acid structure through a biotinylated primer during conventional PCR or RCA). Depending on the purpose of the experiments some further modifications are required after first incubation (see Section 3.3.3). Then the nucleic acid templates are attached to antidigoxigenin-coated beads through digoxigenin–antidigoxigenin interaction. The following protocol describes the sample preparation and data acquisition steps which we use here. 1. Wash the sample channel twice with 100 μL T50 buffer. 2. Inject 1% BSA solution and incubate for 10 min. 3. Wash out the excess BSA with 100 μL T50 buffer twice. 4. Inject 0.2 mg/mL neutravidin solution into the channel. Incubate for 5 min. Then wash out as in Step 3. 5. Depending on the purpose of experiments, such as translocation, unwinding, or diffusion studies, different constructs of nucleic acids

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are incubated in this step. Generally speaking, 100 μL of 250 pM nucleic acid constructs are incubated for 30 min in the channel. After incubation with the nucleic acid constructs, wash out the unbound constructs with two rinses of 100 μL T50 buffer. Take 3 μL of the antidigoxigenin beads solution and mix with 397 μL T50 buffer. Centrifuge at 18,000  g for 5 min at 4°C. Remove the supernatant and resuspend the pellet, diluting 25–100 . Repeat the dilution step once and then sonicate for 10 s. Inject solution with diluted antidigoxigenin beads into sample channel and incubate for 45 min. Wash out the excess nucleic acid constructs with two rinses of 100 μL T50 buffer. Depending on the purpose of experiments, different protein solutions can be added into channel. In general, protein solution is mixed with imaging buffer to 100 μL. The final concentration of protein solution is 2 nM. Here, Cy3-labeled UvrD is the protein we are interested in. When mixing protein solution with imaging buffer, glucose oxidase solution has to be added into the imaging buffer immediately before injecting the imaging buffer into the sample channel to avoid acidification of solution (Shi, Lim, & Ha, 2010). To prevent the evaporation of solution inside the channel, cover the inlet and outlet of the sample channel with immersion oil droplet. Mount the sealed sample chamber on the microscope, using immersion oil to engage the condenser lens with the slide and the objective lens with the coverslip. Use bright field to look at the coverslip surface through a camera and trap a free bead using optical tweezers. After trapping one free bead, align the position of the position-sensitive photodetector by adjusting the manual stages on which the detector is mounted. Next, find and trap a tethered bead whose radius of tethered diffusion corresponds to the length of the DNA constructs. To determine the origin of the trapped tether on the surface and to apply external force accurately in previous step, record forceextension curves through stretching of the nucleic acid-tether in xand y-direction using a computer-controlled stage movement at 25 pN constant force. Apply a desirable force to the tether by moving the piezo-stage, generally in the range of 1 to 30 pN. Use TIRF-based fluorescence

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excitation to image the movement of fluorescent-labeled protein, Cy3labeled UvrD under desirable external force. In the meanwhile, the photons from Cy3 emission are recorded by an EMCCD and the force signal is obtained from QPD signals as a function of time. 3.3.3 Visualizing Initiation Pathways of Helicase Unwinding To observe the initiation pathways of helicase unwinding, a partial duplex DNA construct is used to incubate in the sample chamber as substrates. The partial duplex DNA is synthesized by the following protocol (Lee et al., 2013). 1. 4957-bp dsDNA constructs are synthesized from the sequence (19,360–24,316) of bacteriophage λ DNA by PCR reaction with one biotinylated primer and one unmodified primer. 2. Terminal transferase and digoxigenin-11-ddUTP are added with purified PCR products to attach digoxigenin at the 30 -end. 3. Immobilize the 5-kbp dsDNA (4957 bp + 1 nt) onto the coverslip via neutravidin and biotin interaction, 10 min (Fig. 1A).

Fig. 1 Preparation of partial duplex DNA constructs. (A) Biotinylated dsDNA constructs are produced by PCR with digoxigenin at the 30 -end of the constructs. The constructs are immobilized on the PEG-passivated surface through biotin–neutravidin interaction. (B) T7 exonuclease is injected into the sample chamber, and because of the steric hindrance only one 50 -end strand which does not have a biotin-label can be digested by T7 exonuclease. (C) The final product is a partial duplex DNA with long 30 -ssDNA tails. The length of ssDNA is determined by the incubation time of T7 exonuclease. When the desired length is achieved, the reaction is quenched and T7 exonuclease washed out with T50 buffer.

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4. T7 exonuclease is incubated with dsDNA constructs in the chamber. Since the 50 -end in one strand is protected as a result of surface immobilization through biotin–neutravidin coupling, the 50 -end of the complementary strand is exposed and, therefore, selectively digested by T7 exonuclease (Fig. 1B). 5. A partial duplex with long 30 -ssDNA tail constructs are generated when the reaction has been quenched. The length of the ssDNA strand can be adjusted depending on the experimental purpose by changing the incubation time of the exonuclease reaction (Fig. 1C). After the digestion process of T7 exonuclease, inject and attach the antidigoxigenin beads to the partial duplex DNA substrates. Add imaging buffer and fluorescently labeled helicase into the sample chamber. When applying constant force, in the case of UvrD helicase, one can observe the monomeric form of UvrD helicase translocate and stall at the ss/ds junction (Fig. 2A). It remains stalled until the dissociation of the single UvrD helicase or the formation of a UvrD dimer (Fig. 2B), which initiates the unwinding process (Lee et al., 2013). This assay is extremely useful to examine the stoichiometry of helicases in initiating and maintaining unwinding function in real time. In a different scenario, one can use the assay to examine how cofactors regulate the unwinding processes of helicases. Since one can generate a construct of adequate lengths of single-stranded nucleic acids and

Fig. 2 Schematic illustration of initiation of unwinding pathway. (A) Monomeric form of helicase, in this case UvrD, translocates on ssDNA until it is stalled by a ss/ds DNA junction. Based on a previous report (Lee et al., 2013), the monomeric UvrD stops at the junction. If a second UvrD binds before the first one dissociates, then the unwinding proceed. Otherwise, the first UvrD stays in place until it dissociates from the ss/ds DNA junction site. (B) A dimeric UvrD is sufficient to initiate the unwinding of dsDNA. The real time and synchronized recording of fluorescence intensity and force trajectories allow us to monitor the unwinding rate, stoichiometry of unwinding complexes, and binding time. The arrows show the unwinding direction.

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double-stranded nucleic acids and use optical tweezers as an external force to stretch the substrate linearly, the unwinding dynamics of the complex, helicase, and its cofactor can be monitored in detail. The results of this assay can used to obtain the unwinding rate of helicase, stoichiometry of the helicase complex used to initiate the unwinding, the processivity of helicase unwinding, and the binding time of the complexes, either helicase only or helicasecofactor.

3.3.4 Imaging Helicase Translocation One of the vital functions of helicases besides unwinding is translocation. In this assay, we used secondary-structure-free, long ssNA (see later for the steps of the protocol) as substrates to reveal the kinetic properties of helicases in a minimally interfering way (Fig. 3). With the aid of optical tweezers, long ssNA are stretched out linearly and single-fluorophore localization allows us to monitor the movement of helicase directly in real time. The time trajectories of applied force and the time series of the fluorescence images of the nucleic acid-tether and fluorescently

Fig. 3 Schematic illustration of helicase translocation. To observe the dynamics of helicase translocation on long ssDNA, a long secondary-structure-free ssDNA is immobilized on the PEG-passivated surface and serves as a 1-dimensional platform to observe helicase motions. The advantage of using external tension by optical trapping is not only to maintain the linear form of long ssDNA but also to observe the potential involvement of mechanical force with translocation motion (Lin et al., 2014). The arrows show the translocation direction.

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labeled helicase provides the translocation rate, processivity of helicase translocation, binding time, stoichiometry of translocation complexes, and possible mechanical features generated by helicase translocation. To overcome the challenge of ssDNA forming secondary structures, we use the RCA approach to synthesize a long secondary-structure-free ssDNA substrate for our single-molecule optical tweezers experiments (Fig. 4). With proper design of the template, secondary structures can be avoided in RCA. More importantly, the ssDNA product can be longer than 10 kb, which is difficult to achieve by standard PCR. The following steps refine the synthesis and purification of secondary-structure-free ssDNA, which has been reported previously (Brockman, Kim, & Schroeder, 2011; Lee et al., 2014). 1. 0.2 μM ssDNA template is annealed to 0.2 μM biotinylated primer at 70°C for 2.5 min and then slowly cooled to room temperature. 2. 600 U of T4 DNA ligase are then added to ligate the nick at 16°C for 5 h to form covalently closed circular ssDNA templates. 3. Incubate 50 nM closed circular ssDNA template, 25 μM each of dTTP and dCTP, 20 mg/mL BSA, and 5 U phi29 DNA polymerase together with 1  phi29 DNA polymerase buffer for 12 min (generating >20 kb ssDNA). 4. After the replication process, 2.5 μM digoxigenin-11-ddUTP (Roche) is added to quench the reaction and label the 30 -end of the ssDNA product with digoxigenin. 3.3.5 Mechanical Regulation of Helicase Unwinding/Translocation Finally, we present an assay to study how tensions in nucleic acids substrates regulate the unwinding/translocation of helicase, which has implications for helicase regulation mechanisms. The nucleic acid substrates are the same as in Sections 3.3.3 and 3.3.4. Unlike previous assays which were mentioned in the paragraphs earlier, the piezo-stage is moved to several different positions and remains for a period of time for different constant force measurements. For example, the piezo-stage can be moved from the origin to apply 4, 8, 12, 16, or 20 pN, respectively, and stay in each position for a desired period of time while recording the emitted fluorescence signal from the labeled helicase as a function of time (Fig. 5). Based on the results, one can easily examine how tension affects the functions of helicase.

Fig. 4 Schematic of ssDNA preparation by rolling circle replication. (A) Biotinylated primer is added to template. A partial duplex is formed between the primer and template due to the complementary sequence (red to magenta, green to light green). (B) The nick at the template site is ligated by ligase. (C) phi29 polymerase and dNTP are added into the reaction, and the synthesis of secondary-structure-free, long ssDNA proceeds.

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Fig. 5 Assay to study mechanical regulation of helicase functions. With the control program, the setup is able to record helicase functions as a function of force. The recording time for specific applied forces can be varied for different helicase systems. (A–C) This is an example showing that the translocation of helicase is recorded as a function of force. In this set up, the displacement of the piezo-stage determines the applied force.

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ACKNOWLEDGMENTS These studies were supported by grants from the National Institutes of Health (GM065367) and the National Science Foundation (PHY-1430124) to T.H. We would like to thank Olivia Yang for proofreading the manuscript and Dr. Kyung Suk Lee for constructing the original instrument and training. T.H. is an investigator of the Howard Hughes Medical Institute.

REFERENCES Brockman, C., Kim, S. J., & Schroeder, C. M. (2011). Direct observation of single flexible polymers using single stranded DNA. Soft Matter, 7(18), 8005–8012. http://dx.doi.org/ 10.1039/C1SM05297G. Donmez, I., & Patel, S. S. (2006). Mechanisms of a ring shaped helicase. Nucleic Acids Research, 34(15), 4216–4224. http://dx.doi.org/10.1093/nar/gkl508. Greenleaf, W. J., Woodside, M. T., & Block, S. M. (2007). High-resolution, single-molecule measurements of biomolecular motion. Annual Review of Biophysics and Biomolecular Structure, 36, 171. Ha, T., Enderle, T., Ogletree, D. F., Chemla, D. S., Selvin, P. R., & Weiss, S. (1996). Probing the interaction between two single molecules: Fluorescence resonance energy transfer between a single donor and a single acceptor. Proceedings of the National Academy of Sciences of the United States of America, 93(13), 6264–6268. Jankowsky, E., & Fairman, M. E. (2007). RNA helicases—One fold for many functions. Current Opinion in Structural Biology, 17(3), 316–324. http://dx.doi.org/10.1016/j.sbi.2007. 05.007. Jankowsky, E., Gross, C. H., Shuman, S., & Pyle, A. M. (2001). Active disruption of an RNA-protein interaction by a DExH/D RNA helicase. Science, 291(5501), 121–125. http://dx.doi.org/10.1126/science.291.5501.121. Joo, C., Balci, H., Ishitsuka, Y., Buranachai, C., & Ha, T. (2008). Advances in singlemolecule fluorescence methods for molecular biology. Annual Review of Biochemistry, 77, 51. Lee, K. S. (2013). Fluorescence imaging of single molecule dynamics on long single stranded DNA. Doctor of Philosophy Doctoral dissertation, University of Illinois at UrbanaChampaign. Retrieved from, http://hdl.handle.net/2142/42452. Lee, K. S., Balci, H., Jia, H., Lohman, T. M., & Ha, T. (2013). Direct imaging of single UvrD helicase dynamics on long single-stranded DNA. Nature Communications, 4, 1878. http:// dx.doi.org/10.1038/ncomms2882. Lee, K. S., Marciel, A. B., Kozlov, A. G., Schroeder, C. M., Lohman, T. M., & Ha, T. (2014). Ultrafast redistribution of E. coli SSB along long single-stranded DNA via intersegment transfer. Journal of Molecular Biology, 426(13), 2413–2421. http://dx.doi. org/10.1016/j.jmb.2014.04.023. Lin, C.-T., & Ha, T. (in press). Probing single helicase dynamics on long nucleic acids through force-fluorescence measurement, In A. Gennerich (Ed.), Methods in molecular biology: Vol. 1486 (pp.295–316). Lin, C.-T., Tritschler, F., Suk Lee, K., Gu, M., Rice, C. M., & Ha, T. (2014). Singlemolecule imaging reveals the translocation dynamics of hepatitis C virus NS3 helicase. Biophysical Journal, 106(2), 72a. http://dx.doi.org/10.1016/j.bpj.2013.11.474. Lohman, T. M., & Bjornson, K. P. (1996). Mechanisms of helicase-catalyzed DNA unwinding. Annual Review of Biochemistry, 65(1), 169–214. Lu, H. P., Xun, L., & Xie, X. S. (1998). Single-molecule enzymatic dynamics. Science, 282(5395), 1877–1882. http://dx.doi.org/10.1126/science.282.5395.1877.

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Mackintosh, S. G., & Raney, K. D. (2006). DNA unwinding and protein displacement by superfamily 1 and superfamily 2 helicases. Nucleic Acids Research, 34(15), 4154–4159. http://dx.doi.org/10.1093/nar/gkl501. Martin, R. S., Mark, S. D., & Dale, B. W. (2007). Structure and mechanism of helicases and nucleic acid translocases. Annual Review of Biochemistry, 76(1), 23–50. http://dx.doi.org/ 10.1146/annurev.biochem.76.052305.115300. Matson, S. W., Bean, D. W., & George, J. W. (1994). DNA helicases: Enzymes with essential roles in all aspects of DNA metabolism. BioEssays, 16(1), 13–22. http://dx.doi.org/ 10.1002/bies.950160103. Moffitt, J. R., Chemla, Y. R., Izhaky, D., & Bustamante, C. (2006). Differential detection of dual traps improves the spatial resolution of optical tweezers. Proceedings of the National Academy of Sciences of the United States of America, 103(24), 9006–9011. http://dx.doi. org/10.1073/pnas.0603342103. Robison, A. D., & Finkelstein, I. J. (2014). High-throughput single-molecule studies of protein–DNA interactions. FEBS Letters, 588(19), 3539–3546. http://dx.doi.org/10. 1016/j.febslet.2014.05.021. Roy, R., Hohng, S., & Ha, T. (2008). A practical guide to single molecule FRET. Nature Methods, 5(6), 507–516. http://dx.doi.org/10.1038/nmeth.1208. Schmid, S. R., & Linder, P. (1992). D-E-A-D protein family of putative RNA helicases. Molecular Microbiology, 6(3), 283–292. http://dx.doi.org/10.1111/j.1365-2958.1992. tb01470.x. Selvin, P. R., & Ha, T. (2008). Single-molecule techniques: A laboratory manual. Cold Spring Harbor, NY: Cold Spring Harbor Laboratory Press. Shi, X., Lim, J., & Ha, T. (2010). Acidification of the oxygen scavenging system in singlemolecule fluorescence studies: In Situ sensing with a ratiometric dual-emission probe. Analytical Chemistry, 82(14), 6132–6138. http://dx.doi.org/10.1021/ac1008749. Tuteja, N., & Tuteja, R. (2004). Prokaryotic and eukaryotic DNA helicases. Essential molecular motor proteins for cellular machinery. European Journal of Biochemistry, 271(10), 1835–1848. http://dx.doi.org/10.1111/j.1432-1033.2004.04093.x. Vindigni, A. (2007). Biochemical, biophysical, and proteomic approaches to study DNA helicases. Molecular BioSystems, 3(4), 266–274. http://dx.doi.org/10.1039/B616145F. Visscher, K., Gross, S. P., & Block, S. M. (1996). Construction of multiple-beam optical traps with nanometer-resolution position sensing. IEEE Journal on Selected Topics in Quantum Electronics, 2(4), 1066–1076. http://dx.doi.org/10.1109/2944.577338. Walter, N. G., Huang, C. Y., Manzo, A. J., & Sobhy, M. A. (2008). Do-it-yourself guide: How to use the modern single-molecule toolkit. Nature Methods, 5, 475. Zhou, R., Schlierf, M., & Ha, T. (2010). Chapter sixteen—Force–fluorescence spectroscopy at the single-molecule level. In G. W. Nils (Ed.), Methods in Enzymology: Vol. 475 (pp. 405–426): Cambridge, MA: Academic Press.

CHAPTER SIX

High-Resolution Optical Tweezers Combined With Single-Molecule Confocal Microscopy K.D. Whitley*, M.J. Comstock{, Y.R. Chemla*,†,1 *University of Illinois at Urbana–Champaign, Urbana, IL, United States † Center for the Physics of Living Cells, University of Illinois at Urbana–Champaign, Urbana, IL, United States { Michigan State University, East Lansing, MI, United States 1 Corresponding author: e-mail address: [email protected]

Contents 1. Introduction 2. Optical Trapping and Single-Molecule Fluorescence 2.1 Principles of Optical Trapping 2.2 Single-Molecule Fluorescence Detection 2.3 Combined Optical Tweezers/Single-Molecule Fluorescence Microscope 3. Instrument Design 3.1 Overview 3.2 Interlacing and Time-Sharing 3.3 Optical Layout 3.4 Data Acquisition and Instrument Control 4. Instrument Alignment 4.1 Temporal Alignment 4.2 Co-Alignment of Trapping and Excitation Lasers 5. Combined Optical Trap/smFRET Assay 5.1 Protein Expression and Fluorescent Labeling 5.2 DNA Hairpin Construct 5.3 Sample Flow Chamber 5.4 Sample and Buffer Preparation for Optical Trap–smFRET Assay 5.5 Optical Trap–smFRET Measurement Acknowledgments References

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Abstract We describe the design, construction, and application of an instrument combining dualtrap, high-resolution optical tweezers and a confocal microscope. This hybrid instrument allows nanomechanical manipulation and measurement simultaneously with single-molecule fluorescence detection. We present the general design principles that

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overcome the challenges of maximizing optical trap resolution while maintaining single-molecule fluorescence sensitivity, and provide details on the construction and alignment of the instrument. This powerful new tool is just beginning to be applied to biological problems. We present step-by-step instructions on an application of this technique that highlights the instrument’s capabilities, detecting conformational dynamics in a nucleic acid-processing enzyme.

1. INTRODUCTION Single-molecule techniques have become powerful tools to study fundamental biological processes. Two broadly defined categories are forcebased manipulation and detection techniques (e.g., optical tweezers, magnetic tweezers, AFM, nanopores) and single-molecule fluorescence imaging and spectroscopy. In recent years, a new generation of tools combining both categories has emerged. For example, new hybrid instruments combining optical trapping with single-molecule fluorescence (Bianco et al., 2001; Heller et al., 2013; Hohng et al., 2007; Lang, Fordyce, Engh, Neuman, & Block, 2004; Lee, Balci, Jia, Lohman, & Ha, 2013; van Mameren et al., 2006) have allowed new avenues of investigation, making possible measurement of multiple biomolecular parameters simultaneously. In this chapter, we describe an instrument combining dual-trap optical tweezers with a confocal microscope (Figs. 1 and 2) (Comstock, Ha, & Chemla, 2011). This instrument has the ability to resolve mechanical signals at subnanometer spatial resolution (with the optical traps) and to detect simultaneously the emitted light from a single fluorophore (with the confocal microscope). Applications of this method have just begun to emerge (Comstock et al., 2015; Suksombat, Khafizov, Kozlov, Lohman, & Chemla, 2015), with new results on conformational dynamics of nucleoprotein complexes detected with optical traps and single-molecule F€ orster Resonance Energy Transfer (smFRET). Below, we provide a general overview of optical traps and single-molecule fluorescence, the challenges in combining them, the design principles of our instrument, and its alignment procedures. We end with protocols for replicating a recently reported experiment on the DNA helicase UvrD and the relationship between its conformational state and unwinding activity enabled by this instrument (Comstock et al., 2015).

Fig. 1 Combined high-resolution optical tweezers and confocal microscope. Dual optical traps (outer cones) hold polystyrene microspheres (spheres) tethered by a DNA construct (here a DNA hairpin), while a confocal microscope (middle cone) detects fluorescence from a single molecule. In this example, the conformational and unwinding dynamics of E. coli UvrD helicase are investigated. UvrD helicase exists in two conformational states—“open” (shown in the free protein) and “closed” (shown in the bound protein)—that are differentiated by smFRET between a donor–acceptor pair labeling the protein (green and red disks, respectively). The proteins in this figure were prepared with VMD (Humphrey, Dalke, & Schulten, 1996) from PDB entries 2IS2 and 3LFU. Figure reproduced from Comstock, M. J., Whitley, K. D., Jia, H., Sokoloski, J., Lohman, T. M., Ha, T., & Chemla, Y. R. (2015). Direct observation of structure-function relationship in a nucleic acid-processing enzyme. Science, 348(6232), 352–354 with permission from AAAS.

Fig. 2 Detailed layout of the instrument (not to scale). The instrument consists of three modules: Optical trap (yellow solid lines), confocal excitation (green solid lines) and emission (red dashed lines), and bright-field imaging (blue dotted lines). The asterisk (*) denotes planes conjugate to AOM1, the double cross ({) those conjugate to the steerable mirror (SM). Arrows indicate adjustable translational or rotational stages. Dotted lines indicate the back-focal planes of the objectives. Refer to text for details.

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2. OPTICAL TRAPPING AND SINGLE-MOLECULE FLUORESCENCE 2.1 Principles of Optical Trapping Optical tweezers utilize the momentum carried by light to exert forces on microscopic objects. An infrared (IR) laser tightly focused to a diffractionlimited spot by a high-numerical aperture (NA) microscope objective generates optical forces that can trap a dielectric object—such as a μm-sized polystyrene or glass bead—stably in three dimensions (Ashkin, 1986). Near the focus of light, the optical trap behaves as a linear spring, exerting a force on the trapped object proportional to its displacement. This displacement is typically detected by back-focal-plane interferometry (Gittes & Schmidt, 1998), in which the interference pattern between the incident light and that forward-scattered by the trapped object is imaged onto a position-sensitive photodetector. With proper calibration of the instrument, this signal can be converted into a displacement in nanometers and a force in piconewtons. The sensitivity of optical tweezers has made them a powerful tool to investigate biomolecules at the single-molecule level. By tethering molecules to beads held in traps and applying force, optical tweezers have provided new insights on mechanical, structural, and dynamic properties of biomolecules (Bustamante, Bryant, & Smith, 2003; Heller, Hoekstra, King, Peterman, & Wuite, 2014; Ritchie & Woodside, 2015). They have also been well suited to studying the mechanisms of molecular motors involved in a range of functions—cytoskeletal transport, the central dogma, and beyond (reviewed in Bustamante, Cheng, & Mejia, 2011; Heller et al., 2014; Veigel & Schmidt, 2011). Nucleic acid-processing motors in particular are studied by monitoring the extension of the DNA or RNA molecules tethered by the trapped beads (for example, Fig. 1). These molecular tethers often serve an additional role to position the systems of interest away from the high light intensity of the optical traps. Advances in instrument design over the last dozen years have increased optical tweezers sensitivity remarkably. Instruments with active stage stabilization (Carter et al., 2007) and others incorporating dual traps formed from the same laser (Abbondanzieri, Greenleaf, Shaevitz, Landick, & Block, 2005; Moffitt, Chemla, Izhaky, & Bustamante, 2006) have improved instrument stability to such an extent that it is now possible to detect subnanometer signals. These so-called high-resolution optical

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tweezers have enabled investigations of nucleic acid-processing molecular motors in extreme detail (Abbondanzieri et al., 2005; Cheng, Arunajadai, Moffitt, Tinoco, & Bustamante, 2011; Moffitt et al., 2009; Qi, Pugh, Spies, & Chemla, 2013).

2.2 Single-Molecule Fluorescence Detection Another powerful technique to study biological processes relies on the detection of light emitted by individual fluorophores (Moerner, 2007). Labeling proteins, nucleic acids, or small molecules with a fluorophore— commonly, organic dyes such as Cy3, Cy5, and TMR—allows measuring their spatiotemporal dynamics at the single-molecule level. Single-molecule fluorescence detection can be incorporated into the standard types of optical microscopy—bright field, confocal, or total internal reflection (TIR)— requiring only an excitation light source at a wavelength tuned to the absorption spectrum of the fluorophore and a sensitive charged-coupled device (CCD) camera or avalanche photodiode (APD) to detect the emitted light efficiently. Fluorescence provides several measurable quantities that can provide information on biomolecular conformational states and their dynamics: intensity, lifetime, position, and orientation. These can be used for tracking diffusion or directed motion of a labeled molecule, to detect the binding of labeled molecules to a system of interest, or to determine the stoichiometry or composition of multicomponent complexes (Joo, Balci, Ishitsuka, Buranachai, & Ha, 2008). A powerful tool for probing conformational dynamics of biomolecules is smFRET. Here, a “donor” molecule excited to a high-energy state transfers its energy to a neighboring “acceptor” molecule via induced-dipole interactions, which then emits light of a longer wavelength. The efficiency of energy transfer (or FRET efficiency), E, is strongly dependent on the distance R between the donor and acceptor,   varying as E ¼ 1= 1 + ðR=R0 Þ6 , where the F€ orster radius R0 is the distance ˚ for the Cy3-Cy5 pair). at which 50% of the energy is transferred (R0 ¼ 60 A Thus, smFRET is a spectroscopic technique that measures the distance between a single donor–acceptor fluorescent dye pair (Ha, 2001; Ha et al., 1996) and is sensitive to conformational changes typically in the ˚ distance range (Forster, 1965; Stryer & Haugland, 1967). 30–80 A This powerful technique has been used to study a wide variety of biological systems (Kim & Ha, 2013).

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2.3 Combined Optical Tweezers/Single-Molecule Fluorescence Microscope In recent years, a number of hybrid instruments combining optical tweezers with fluorescence detection have been developed. This work has been motivated by the desire to detect biomolecular dynamics with multiple orthogonal probes simultaneously. Optical tweezers have been combined with many standard forms of fluorescence microscopy—bright-field (Bianco et al., 2001; van Mameren et al., 2006), TIRF (Lang et al., 2004; Lee et al., 2013), confocal (Hohng et al., 2007), and even STED microscopy (Heller et al., 2013). Example applications of these new techniques include the tracking of singly labeled biomolecules on DNA stretched with optical traps, imaging of nucleoprotein complexes on extended DNA, and measurement of biomolecular conformational dynamics by smFRET as a function of force (reviewed in Chemla, 2016; Heller et al., 2014). The ability to measure and/or control multiple variables simultaneously has provided many new insights not available when using each method independently. An important technical challenge to combining optical traps with singlemolecule fluorescence detection is that many fluorophores photobleach very rapidly (1–2 s) when located in the light field of the optical trap (van Dijk, Kapitein, van Mameren, Schmidt, & Peterman, 2004). This phenomenon results from a two-photon process in which a fluorophore absorbs a fluorescence excitation photon to its excited state followed by an IR optical trap photon to a higher energy level. Rather than decaying to the ground state and emitting light, the fluorophore instead decays to an ionized dark state. In the applications mentioned above, this problem was circumvented by imaging fluorescence far from the optical trap light field. This approach requires using long, compliant molecules to separate the trap(s) from the fluorescence imaging region spatially. Unfortunately, this reduces instrument resolution, which is proportional to the stiffness of the molecular tether (Moffitt et al., 2006). An alternative is to strobe the two light sources out of phase, separating them temporally (Brau, Tarsa, Ferrer, Lee, & Lang, 2006). As shown by Brau et al., interlacing trapping and fluorescence excitation laser significantly increases the length of time before fluorophores photobleach, allowing spatially overlapping light sources. Recently, we integrated these advances to develop an instrument combining high-resolution optical traps with a single-molecule fluorescence microscope (Comstock et al., 2011). The sensitivity of this instrument allows detection of fluorescence signals with single-molecule sensitivity

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simultaneously with mechanical displacements at sub-nm resolution. This approach is starting to be adopted by other groups (Duesterberg, FischerHwang, Perez, Hogan, & Block, 2015; Sirinakis, Ren, Gao, Xi, & Zhang, 2012) and has begun to provide new insights into biomolecular systems (Comstock et al., 2015; Suksombat et al., 2015). In the following sections, we describe the design principles and construction of this hybrid instrument. We also provide detailed protocols for its application, recapitulating measurements of conformational dynamics in a nucleic acidprocessing enzyme (Fig. 1) (Comstock et al., 2015).

3. INSTRUMENT DESIGN 3.1 Overview In this section we provide a general overview of the instrument design. Due to their exceptional stability, we utilize dual-trap optical tweezers in which both traps are formed from the same IR laser (Bustamante, Chemla, & Moffitt, 2008; Moffitt et al., 2006). In the dual-trap design, the traps are formed microns from the sample chamber surface. As a result, TIR microscopy, in which fluorescence excitation occurs only in the exponentially decaying evanescent field 100 nm off the surface, is not well suited for this configuration. In our design, we use confocal microscopy instead, where we position the confocal excitation spot in the same plane as the two trapped beads (Fig. 1). High-resolution measurements have so far been made using short (1 μm), stiff tethers (e.g., Abbondanzieri et al., 2005; Cheng et al., 2011; Moffitt et al., 2009; Qi et al., 2013). In such a configuration, the optical traps and confocal excitation spot occupy the same volume and the light sources overlap significantly. To avoid fast photobleaching of fluorophores by absorption of IR and visible photons, we interlace the two light sources. Thus, the instrument incorporates and adapts design elements of previous setups: (1) dual-trap high-resolution optical tweezers (Moffitt et al., 2006), (2) combined optical trap/confocal microscope (Hohng et al., 2007), and (3) interlaced trap and fluorescence excitation light sources (Brau et al., 2006).

3.2 Interlacing and Time-Sharing 3.2.1 Interlacing With Acousto-Optic Modulators A critical component of the instrument design is the interlacing of the trapping and fluorescence excitation light sources. As shown in previous work

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(Brau et al., 2006), the interlacing rate is an important consideration. If the rate is too low, the trap stiffness will be lower than that expected from the average trapping laser power (i.e., peak power  interlacing duty ratio) because the trapped bead moves beyond the trapping region when the beam is turned OFF for long durations. (A free 1-μm-diameter bead experiencing a 1 pN force will move by 100 nm in 1 ms.) Thus, it is critical to modulate the trapping beam at a sufficiently high frequency fm. A good practice is to operate above the characteristic frequency of the bead in the trap, fc ¼ κ/2πγ, where κ is the stiffness and γ ¼ 6πηr is the Stokes drag of the bead (η is the viscosity and r is the bead radius). Typically fm needs to be >10 kHz. These interlacing frequencies are feasible with acousto-optic devices. In an acousto-optic modulator (AOM), a radio frequency (RF) sound wave generated by a piezoelement propagates inside a crystal. An incident beam passing through this crystal is diffracted by the periodic modulation of the index of refraction generated by the sound wave. An AOM behaves essentially as a tunable diffraction grating, where the sound wave amplitude determines the intensity of the diffracted laser beam and its frequency the diffraction angle. The maximum frequency at which the AOM can modulate a laser beam is approximately given by the speed of sound in the crystal divided by the beam diameter. In our instrument, a 1.5-mm beam waist corresponds to 2 MHz, and we interlace at a frequency of fm ¼ 66 kHz, well above fc. Two separate AOMs modulate the trapping (Fig. 2, AOM1; IntraAction, ATM-803DA6B) and fluorescence excitation beams (AOM2; IntraAction, AOM-802AF1) out of phase. This requires synchronizing the interlacing cycles of both AOMs with μs-level precision, achieved through a data acquisition and instrument control architecture that we discuss below (see Section 3.4). The RF source that drives the AOM sound wave is an important component of the setup, as precise and stable RF generation is critical for stable trap positioning. We chose to build a custom RF synthesizer rather than use an integrated commercial source, as the former exhibits significantly lower noise (Comstock et al., 2011). This RF source consists of a temperaturecompensated crystal oscillator (TCXO; Conner-Winfield, HTFL5FG5049.152M) mounted on a direct digital synthesis RF synthesizer board (Analog Devices, AD9852/PCBZ or AD9854/PCBZ). The TCXO provides a 49.152 MHz fixed output reference frequency with 1 ppm stability. The stability of this clock frequency has a direct effect on the stability of the RF output frequency and the stability of the trap positions. We configure the RF synthesizer board to multiply the clock frequency 6  to a final value

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of  300 MHz. Finally, a low-noise, fixed-gain RF power amplifier (Mini-Circuits, ZHL-5 W-1) amplifies the synthesizer output signal to drive the AOM. A commercial RF source (IntraAction, ME-801.5-6) is used to drive the AOM modulating the fluorescence excitation, since an ultra-stable RF frequency is not as critical in this case. 3.2.2 Formation of Dual Traps by Time-Sharing In many dual-trap instruments, the two traps are generated by separating the trapping laser into orthogonally polarized beams. One of the two beams is then independently controlled by an actuated tip–tilt mirror to displace one trap relative to the other in the specimen plane (Bustamante et al., 2008; Moffitt et al., 2006). In our instrument, we use the interlacing AOM both to generate and steer the traps by controlling the beam diffraction angle with the RF drive frequency. We exploit the high speed of acousto-optic devices to generate the dual traps by time-sharing, deflecting the trapping beam between two angles (corresponding to two positions in the specimen plane) at fast rates (Visscher, Brakenhoff, & Krol, 1993; Visscher, Gross, & Block, 1996). Although acousto-optic deflectors (AODs) are optimized to provide a larger deflection range and are more commonly used for steering than AOMs, we found that they affect beam shape and quality. In addition, we have observed that beams deflected by AODs exhibit larger fluctuations in power over small angles than those deflected by AOMs. We thus use an AOM instead of an AOD despite the smaller deflection range. Time-sharing the two traps has advantages over more common polarization-based designs. When splitting the trapping light by polarization, the orthogonally polarized beams travel along separate paths and must pass through different optical components in order to be independently controllable. Prior studies (Bustamante et al., 2008) demonstrated that the longer this differential optical path is, the more susceptible the instrument can be to environmental noise, because each beam is subject to different local environments. In contrast, our approach keeps the differential path between traps to an absolute minimum, since the time-shared beams share identical optical components and a virtually identical beam path. Another issue affecting polarization-based designs is that microscope objectives mix polarization, which leads to cross-talk and interference between the two traps that is separation dependent (Bustamante et al., 2008). These artifacts are eliminated in the time-sharing approach. In summary, we use AOM1 and AOM2 to interlace the trapping and fluorescence excitation lasers, respectively, where AOM1 is also responsible

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Fig. 3 Interlacing and time-sharing timing. Trap 1 and trap 2 are ON each for 1/3 of the cycle and fluorescence excitation is ON during the final 1/3. Plotted are the laser intensities measured by feedback photodetectors QPD1 (traps 1 and 2; gray, bottom panel) and PD (fluorescence excitation; black, top panel). Traps 1 and 2 are set to different intensities for clarity. 625-ns delays (shaded regions) between turning OFF (ON) the optical traps and turning ON (OFF) the fluorescence excitation ensure no overlaps between light sources. Trap data acquisition occurs at the time points “x” and “+” for trap 1 and 2, respectively. A digital pulse (black, bottom panel) synchronous with trap data acquisition is output from the DAQ card to make temporal adjustments during alignment. The rising edge of a digital pulse is synchronous with the trap data acquisition (vertical arrows). A digital pulse (dark gray, top panel) synchronous with the APD fluorescence data acquisition is also output from the DAQ card. Emission photons are counted only during this time interval (horizontal arrow). Figure adapted from Comstock, M. J., Ha, T., & Chemla, Y. R. (2011). Ultrahigh-resolution optical trap with single-fluorophore sensitivity. Nature Methods, 8(4), 335–340 with permission from Nature Publishing Group.

for both time-sharing and steering the traps. During one 15-μs cycle (1/66 kHz), each trap is turned ON for 1/3 of the cycle, and then both remain OFF for the final 1/3 cycle when the fluorescence excitation is ON (Fig. 3).

3.3 Optical Layout In the following section we describe the overall optical layout. The instrument is organized around three “modules” described below: (1) dual-trap optical tweezers (Fig. 2; yellow lines), (2) a fluorescence confocal microscope (green lines for excitation, red and pink dashed lines for emission),

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and (3) a bright-field imaging system (blue lines). Each module has its own illumination source and detectors, and is partitioned with blackout enclosures to minimize stray light. Please note that some details on the assembly of this instrument are omitted, as optical trap designs and construction protocols have been discussed extensively in the literature (Block, 1998; Bustamante et al., 2008; van Mameren, Wuite, & Heller, 2011). 3.3.1 Trapping Module A single-mode, polarization-maintaining fiber laser (IPG Photonics, YLR5-1064-LP; λ ¼ 1064 nm, 5 W) generates the trapping beam. This type of laser provides an ideal Gaussian mode and excellent pointing stability. After an optical isolator (ISO1), we make coarse adjustments to the trapping beam power by passing the laser through a rotary half-wave plate (HW1) and a polarizing beam splitter (PBS1) cube, diverting a fraction of the power into a high-power beam dump (BD). The trapping beam next passes through AOM1. The beam entering the AOM has a diameter smaller than the 3-mm “active area” of the sound field in this device (1.5 mm) to maximize the modulation rate of the AOM. The first-order diffraction beam out of the AOM is next collimated and expanded in two stages by telescope T1 (expanding the beam by 3 ) and T2 (by 2 ) to a final beam diameter of 9 mm, overfilling the back aperture of the front objective (8 mm) (Neuman & Block, 2004). The telescopes also serve as imaging systems that make the pivot plane for the trapping beam inside the AOM conjugate to the back-focal plane of the objective (denoted by * in Fig. 2). Thus, deflecting the trapping beam with the AOM displaces the traps in the sample plane without clipping the beam on the objective aperture. In practice, there is no single pivot plane inside the AOM about which the trapping beam rotates, so this can only be achieved approximately. Between the two telescopes, we deflect a small percentage (1–2%) of the trapping beam with a high-quality wedged beam sampler (BS1; Newport, 10Q20NC.3) to a power feedback stage. In all acousto-optic devices, the intensity of the diffracted beam varies as a function of deflection angle, resulting in a trap stiffness that changes as the trap is steered in the sample plane. The power feedback stage eliminates these unwanted changes in the trapping beam intensity. Measuring the intensity of the sampled beam with a photodetector (QPD1; First Sensor, QP154-Q-HVSD), the instrument control system (see Section 3.4) modulates the RF amplitude driving AOM1 to ensure a constant trapping beam intensity. This power feedback

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stage reduces intensity noise in the trapping beam by up to 6 orders of magnitude across a broad frequency band (Comstock et al., 2011). We next reflect the trapping beam into the front objective (O1) using a short-pass dichroic mirror (D1; reflected wavelength ¼ 1064 nm, transmitted wavelength ¼ 415–700 nm). Similar to a prior high-resolution optical tweezers design (Bustamante et al., 2008; Moffitt et al., 2006), we use waterimmersion objectives (Nikon 1.2 NA, CFI Plan APO VC 60XWI). Although they generate slightly weaker traps and have a lower fluorescence collection efficiency than oil-immersion objectives due to their smaller NA, water-immersion objectives have the advantage of focusing independently of sample depth. Thus, the optical traps are better decoupled from surface drift, improving stability (Bustamante et al., 2008). O1 forms the optical traps inside the sample chamber by focusing the trapping beam, and an identical back objective (O2) collects the trapping light. The sample chamber is mounted on a three-axis motorized translation stage (Newport, Ultralign 562-XYZ) controlled by motorized actuators (Newport, ESP301-3 N and TRA12CC), so the traps can be moved to different areas of the sample (see Section 5.5). A second dichroic (D2) reflects the trap light collected by O2 into the detection stage, where the trapped bead positions are monitored by a quadrant photodetector (QPD2; First Sensor, QP154-Q-HVSD). Both QPD1 and QPD2 are IR-enhanced photodetectors and do not suffer from the parasitic, low-pass (10 kHz) filtering exhibited at 1064 nm by more conventional silicon-based position-sensitive detectors (PSDs) (Huisstede, van Rooijen, van der Werf, Bennink, & Subramaniam, 2006). This choice of photodetector is particularly important because of the fast interlacing and time-sharing frequency, 66 kHz. Note that a single detector, QPD2, monitors the positions of both trapped beads. Since the traps are time-shared, a separate signal from each bead is obtained during different phases of the cycle (Fig. 3). Thus, data acquisition must be tightly synchronized with the timesharing/interlacing cycle so that the appropriate signal is recorded at the correct time (see Section 4.1). 3.3.2 Fluorescence Confocal Module We next describe the layout for the fluorescence microscope module. A 532-nm excitation laser beam (50 mW; Spectra-Physics Excelsior, 53250-CDRH) passes through an optical isolator, a power stage, an AOM, a power stabilization stage, and two beam expansion stages (Fig. 2). We remind the reader that in contrast to the trap module, the AOM used here

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(AOM2; IntraAction, AOM-802AF1) only controls the intensity of the fluorescence excitation, not its deflection angle. We instead use a tip–tilt steerable mirror (SM; Mad City Labs, Nano-MTA2 Invar) to move this beam laterally (x–y) in the sample plane. The fluorescence excitation beam is collimated and expanded in two stages by telescope T3 (expanding the beam by 4 ) and T4 (by 1.3 ) to a final beam diameter of 6 mm. We underfill the back aperture of the front objective (O1) slightly as this facilitates coalignment with the trapping beam. A short-pass, 532-nm dichroic mirror (CVI, SWP-43-RU532-TUVIS-PW-1025-C or Semrock, FF875Di01-25) reflects the excitation beam into O1, which focuses it into the sample plane. During measurements, the position of the confocal excitation spot is adjusted with SM to match those of the trapped beads in the sample plane. Telescope T4 images the pivot plane of SM at the back-focal plane of O1 (denoted by { in Fig. 2), such that lateral adjustments can be made without clipping the beam. We adjust the axial (z) position of the confocal excitation spot by two translation stages on the first and second lenses of T3 and T4, providing coarse and fine adjustments, respectively. Each stage moves its lens along the optical axis, changing the beam collimation. It is important to note that chromatic aberrations in O1 mean that collimated trapping and fluorescence beams will not be focused in the same plane. Therefore, axial adjustment of the fluorescence excitation is essential for proper alignment (see Section 4.2). The fluorescence emitted at the confocal excitation spot is collected by O1 and travels back through T4. A long-pass dichroic mirror (D3; Semrock, LPD02-532RU-25) between T3 and T4 diverts the emission light into a confocal pinhole stage where out-of-focus light is rejected. All the lenses that comprise the fluorescence emission pathway are achromatic doublet lenses (430–700 nm antireflection coated) to minimize chromatic aberrations across the range of fluorescence excitation and emission wavelengths. The first lens of telescope T5 focuses the emitted light into the pinhole (20-μm diameter) and the second lens collects the transmitted light and focuses it onto two APDs (Excelitas, formerly PerkinElmer, SPCM-AQRH-14). A 590-nm edge dichroic mirror (D6; Chroma, 590dcxr) splits the emission into a donor channel (APD1) and an acceptor channel (APD2) for smFRET. We use a notch filter (F1; Chroma, HQ545lp) to cut out 532-nm excitation light from the confocal pinhole stage, and emission filters (F2 and F3; Chroma, HQ580/60m and HQ680/60m) for the donor and acceptor channels, respectively. For alignment purposes, we also collect the fluorescence

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excitation beam passing through the sample chamber with O2 and divert this light onto a standard silicon-based PSD (First Sensor, DL100-7-PCBA3). 3.3.3 Bright-Field Imaging The last module consists of a video imaging system to visualize the sample chamber during measurements and to image the optical traps and the fluorescence excitation during alignment. The sample plane is illuminated by a blue LED using K€ ohler illumination and imaged onto an IR-enhanced CCD camera (Watec, WAT-902-B). Band-pass filters on motorized flip mounts remove the 1064-nm trapping light (F4, 1000-nm short-pass filter, Thorlabs, FES1000; Fig. 2) and 532-nm fluorescence excitation light (F5, 532-nm single-notch filter, Semrock NF01-532U-25) from the video image during typical measurements. The video signal is recorded and displayed on the instrument computer using a PCI-express video framegrabbing card (Matrix Vision, mvDELTAe-BNC).

3.4 Data Acquisition and Instrument Control The interlacing and time-sharing scheme of the instrument requires coordination between components that control the trapping and fluorescence excitation beams—AOM1 and AOM2—and detectors that collect data— feedback detectors QPD1 and PD, bead position detectors QPD2 and PSD, and single-photon-counting detectors APD1 and APD2. We use a multifunction Field Programmable Gate Array (FPGA) PC data acquisition (DAQ) card (National Instruments, NI PCIe-7852R) to control these components with 25 ns-level synchrony. FPGA-based DAQ cards have an advantage over traditional cards because all key timing operations are programmed directly onto a chip on the card itself. This DAQ architecture avoids any operating system interference from the PC and runs consistently with 40 MHz (25 ns) timing resolution. With the exception of the bright-field imaging system camera signal, all signals are acquired through the FPGA DAQ card (Fig. 4). Eight analog signals are sampled by the card: three inputs from the trap QPD2 monitoring the bead positions (x, y, and sum voltage), three inputs from the fluorescence PSD monitoring the fluorescence excitation beam (x, y, and sum voltage), and one input each for the trap and fluorescence feedback detectors QPD1 and PD monitoring beam intensity (sum voltage). Data from these inputs are acquired in sequence and transferred from the FPGA to the host PC memory using Direct Memory Access (DMA) every 5 μs (1/3 cycle). The DAQ card also has two digital inputs that collect single-photon digital

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Fig. 4 Input/output architecture of the FPGA-based DAQ card. The DAQ card receives analog inputs (AI) from four photodetectors (QPD1, QPD2, PSD, PD; see Fig. 2) and digital inputs (DI) from two avalanche photodiodes (APD1, APD2). It communicates via digital output (DO) lines to the RF synthesizer board, the output of which is sent to an amplifier and then to the trap AOM (AOM1). “Debugging” DO lines are used for synchronizing detection input timing with the interlacing cycle (see Fig. 3). The DAQ card also uses analog output (AO) lines to control the steerable mirror (SM) for the fluorescence excitation. To control the commercial RF synthesizer that drives the fluorescence AOM (AOM2), a DO line from the DAQ card is used along with an AO line from an external high-current analog output card.

pulse trains generated by APD1 and APD2. These signals are counted, integrated, and transferred via DMA (typically every 1 ms). The FPGA outputs are predominantly dedicated to communication with AOM1 and AOM2. (The exceptions are the analog outputs that drive the piezoactuated stage (SM) that steers the confocal excitation in the sample plane.) We use FPGA digital output channels to communicate with the custom-built RF synthesizer driving AOM1 that modulates the optical trap. Every cycle, the FPGA writes RF frequency and amplitude output values to the RF synthesizer chip buffered memory through these digital lines. There are many instrument-specific subtleties in communicating with the RF synthesizer, and detailed protocols for doing so are now in the literature (Whitley, Comstock, & Chemla, 2017). Control of AOM2 that modulates fluorescence excitation is simpler due to the commercial driver for this AOM: an analog output channel controls the beam intensity,

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and a digital output gates the RF. We note that this commercial driver requires an external high-current analog output card (Fig. 4; National Instruments, NI 9265), as the DAQ card cannot provide sufficient current to drive it. This input/output architecture allows feedback loops to be run directly on the FPGA at 66 kHz: trap and fluorescence laser power stabilization and force clamping during measurements.

4. INSTRUMENT ALIGNMENT 4.1 Temporal Alignment Since switching between the two traps and fluorescence excitation occurs at high rates (66 kHz), several components must be carefully synchronized for optimal functioning. Timing considerations generally fall into two categories: synchronization between light sources and synchronization between data acquisition and light source. The former category includes (1) modulating the traps and fluorescence excitation out of phase with each other, and (2) synchronizing changes in trapping beam position and intensity during time-sharing. In the latter category, (1) trapped beam signals acquired by the QPDs must coincide with the appropriate trap ON interval; (2) fluorescence excitation measurements by the feedback PD must occur during the fluorescence ON interval; and (3) fluorescence emission photons must be collected by the APDs in the appropriate ON interval. In this section we describe these timing adjustments for optimal performance. The adjustments performed here are robust, and only need to be made when the instrument is first set up. We make adjustments by viewing signals from various detectors directly on an oscilloscope with the appropriate bandwidth (e.g., a Tektronix MDO4000B, which is a combined DPO oscilloscope and RF spectrum analyzer). In addition, we configure the FPGA with several “debugging” DO lines that output digital pulses synchronous with the data acquisition timing and view these on the oscilloscope (Fig. 3). Detailed protocols are provided elsewhere (Whitley et al., 2017). 4.1.1 Interlacing the Trapping and Excitation Lasers We align the trapping laser intervals with the excitation laser interval by viewing the total intensity signals (sum voltages) from the feedback QPD1 for the trapping beam and PD for the fluorescence beam. The phase of the fluorescence excitation AOM2 is then adjusted programmatically until the fluorescence ON interval is positioned between trap ON intervals

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(Fig. 3). We set additional 625-ns delays between turning OFF (ON) the optical traps and turning ON (OFF) the fluorescence excitation to avoid any overlap between light sources (Brau et al., 2006) (Fig. 3, gray-shaded regions).

4.1.2 Synchronizing Beam Position and Intensity Modulation During Time-Sharing The position and intensity of the trapping laser during time-sharing are controlled by the frequency and amplitude of the RF signal sent to the trap AOM. We find there is a delay (typically 125–250 ns) between switches in beam position and intensity when the FPGA is programmed to switch both simultaneously. Such a delay results in an unwanted sudden change in one trap stiffness. In order to prevent this artifact, we configure the FPGA so that the timing for RF frequency and amplitude changes are controlled independently, and adjust the delay between trigger signals to make the switches synchronous by viewing the RF signal directly on an oscilloscope (Whitley et al., 2017).

4.1.3 Synchronizing Data Acquisition and Interlacing: Trapping Laser Measurements of trapping beam intensity and bead position must be synchronized with the corresponding trap ON interval. Moreover, it is important to realize that each trapped bead moves toward its trap center during the ON interval and away from it during the OFF interval due to the imbalances in force during interlacing. This bead motion can span several nanometers at a 66 kHz interlacing rate, depending on the force (Fig. 5). Since we care about measuring the average force on the bead, position data must be acquired precisely at the time point of each ON interval that corresponds to the average bead position. For interlacing rates fm much faster than the characteristic frequency fc of the bead, the bead displacement is close to linear in time and the average bead position corresponds to the center of the ON interval. To synchronize the trap intervals with the input timing of the DAQ card, we output from the DAQ card a digital pulse synchronous with the analog input timing (Fig. 3, black trace, bottom panel) and view this pulse alongside the trapping laser intensity measured by the feedback QPD1. We adjust the phase of the input timing programmatically until it is centered on each trapping laser ON interval.

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Fig. 5 Bead motion during time-sharing period. Measurements of trapped beads’ positions during one time-sharing–interlacing cycle for three different time-sharing– interlacing rates (16, 33, 66 kHz). The time axis is normalized to 2/3 of the interlacing cycle duration, i.e., the time-sharing period (see Fig. 3) where trap 1 is ON for half and trap 2 is ON for the second half. Bead oscillations due to imbalanced forces (schematic) are minimized by faster time-sharing–interlacing rates. Data acquired in the middle of each trap ON period (points B and B0 ) give the correct average displacement and force, compared to data before (A and A0 ) or after (C and C0 ). Figure reproduced from Comstock, M. J., Ha, T., & Chemla, Y. R. (2011). Ultrahigh-resolution optical trap with single-fluorophore sensitivity. Nature Methods, 8(4), 335–340 with permission from Nature Publishing Group.

4.1.4 Synchronizing Data Acquisition and Interlacing: Fluorescence Excitation As with the trapping laser, detection of the excitation laser by its feedback PD must be synchronized with its ON interval. Due to time delays in the detector analog input, the DAQ card may not sample the excitation laser intensity at the appropriate time even though the two lasers are synchronized. The procedure for synchronizing the excitation laser to its detectors is identical to that for the trapping laser, where a digital pulse synchronous with the analog input timing of the excitation laser detector is viewed

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alongside the excitation laser intensity signal from PD, and then adjusted until the digital pulse is centered on the laser’s ON interval. 4.1.5 Synchronizing Data Acquisition and Interlacing: Fluorescence Emission A final temporal alignment is synchronizing the excitation laser ON interval with fluorescence emission detection by the APDs. The best way we have found to do this is to leave the APDs ON continuously and program the FPGA to record photon counts only during the excitation laser ON interval. As above, synchronizing the FPGA recording interval with the excitation laser ON interval is eased by creating a digital pulse synchronous with the APD measurement ON time and viewing it (Fig. 3, dark gray trace, top panel) alongside the excitation laser ON interval. We typically make this timing adjustment by trapping a fluorescent bead (e.g., 1-μm, 575-nm emission bead; Thermo-Fisher, F-8819) and centering the confocal excitation spot on it (see Section 4.2). The APD measurement ON time is then adjusted until the observed fluorescence emission signal just begins to increase. From this time point, we shift the phase of the APD recording interval forward by 25% of the interlacing cycle. We use the increase of the fluorescence signal for this alignment rather than its decay, because the beads exhibit phosphorescence, leading to a fluorescence signal that does not decay instantly when the excitation laser is turned OFF.

4.2 Co-Alignment of Trapping and Excitation Lasers Procedures for aligning optical tweezers and confocal microscopes have been described in detail elsewhere, and will not be duplicated here. Instead we describe coaligning the optical traps and confocal excitation spot both laterally and axially to the same depth as the trapped beads. As mentioned above, the instrument objectives are not perfectly achromatic, focusing a collimated IR trapping beam at a different depth than a collimated fluorescence excitation beam. The coalignment of the two lasers is done in several stages. A first, coarse alignment is done by imaging the beam spots onto the CCD camera of the bright-field imaging system (Fig. 2). With both beams well-collimated and centered on the back aperture of O1, we adjust the front objective stage until the trapping laser and excitation laser beam spots are both visible on the CCD camera. It is necessary to adjust the intensities of the lasers to ease

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visualization. The traps should appear as two focused spots on the CCD image. We then align the fluorescence excitation beam spot’s lateral (x–y) and axial (z) position to match those of the traps by manually adjusting the rotary stage beneath the SM and the translational stage on the first lens of telescope T3, respectively. Fine adjustments to the x–y and z positions of the fluorescence excitation are made using the piezomotors of SM and the second, movable lens in telescope T4, respectively. Although the axial alignment only needs to be done once, we find that the lateral alignment must be repeated at the start of each experiment. We have devised two procedures for fine lateral and axial alignment: (1) taking an image of a trapped fluorescent bead by confocal raster scan, or (2) using the fluorescence excitation laser as a “detection beam” (Neuman & Block, 2004) for a trapped nonfluorescent bead. In the first method, a fluorescent bead is trapped and its fluorescence signal from APD1 is collected. By scanning the confocal spot with SM over the trapped bead in the x and y directions (Fig. 6A), an image of the bead is taken, from which an optimal lateral alignment can be obtained. For axial alignment, the confocal spot lateral position is set to the center of the bead, and the fine adjustment telescope lens stage position is adjusted incrementally while recording the fluorescence intensity at each position (Fig. 6B). The optimal axial position is the one that maximizes the fluorescence signal. In the second method, we use the fluorescence excitation laser as a detection beam, measuring the position of a trapped nonfluorescent bead (e.g., 810-nm streptavidin bead; Spherotech, SVP-08-10) by back-focal plane interferometry with the PSD. Here it is convenient to use a high beam intensity (3 mW) for a larger signal, although the APDs must be turned off to prevent damage. Lateral alignment is achieved by measuring the beam deflection as the confocal spot is scanned through the trapped bead in x and y using SM (Fig. 6A). For axial alignment, the confocal spot lateral position is set to the center of the bead and the Brownian motion of the bead is measured, similar to when the trap is calibrated (Berg-Sørensen & Flyvbjerg, 2004; Neuman & Block, 2004). By fitting the power spectral density of the bead position, the conversion factors αx and αy between bead position (in nm) and PSD output (in V) are obtained. After plotting αx and αy vs stage position (Fig. 6B), the optimal position is found from the minimum in αx and αy. This position corresponds to the highest detection sensitivity, where the confocal spot is aligned in the same plane as the trapped bead.

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Fig. 6 Spatial alignment of confocal excitation spot and optical traps. Two methods of alignment are shown. (A) Top panel: Lateral alignment by scanning the confocal spot (using SM) over two trapped beads. Signals resulting from bead fluorescence are recorded by APD1 (yellow: higher fluorescence intensity). Middle and bottom panels: Lateral alignment by scanning confocal spot laser over two trapped beads. Here, the PSD measures the deflection in x (middle) and y (bottom) of the laser as it is scanned [black (yellow) denotes negative (positive) deflections]. (B) Axial alignment by two methods. Circles: Axial alignment by scanning the focal depth of the confocal spot (using the translational stage in T4) over a trapped fluorescent bead. The fluorescence intensity measured by APD1 is maximized when the spot is aligned in the plane of the trapped bead. Crosses and exes: Axial alignment by scanning the confocal spot depth over a trapped bead. Here, the PSD voltage-to-bead position conversion factors, α, derived from trap calibration are minimized when the spot is aligned in the plane of the trapped bead.

5. COMBINED OPTICAL TRAP/smFRET ASSAY In the following section, we describe an application of this instrument combining optical trapping and smFRET. The protocols below recapitulate an experiment carried out by Comstock et al. investigating the structure– function relationship of the E. coli DNA helicase UvrD (Comstock et al., 2015). UvrD is involved primarily in DNA repair and utilizes the energy of ATP hydrolysis to translocate on single-stranded DNA in a 30 –50 direction and to separate the strands of the duplex. It is a prototype for the structural superfamily 1 (SF1) of helicases. Crystal structures of UvrD and other SF1 helicases reveal two distinct conformational states, termed “closed” and

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“open” (Fig. 1), where one of its four domains is rotated 160 degrees with respect to the other three (Jia et al., 2011; Lee & Yang, 2006). These two states were long proposed to regulate protein activity, but their exact roles remained highly debated. To understand the relationship between these two structures and UvrD function, Comstock et al. used the hybrid instrument described here to correlate the open/closed conformation of fluorescently labeled protein (measured by smFRET) with its helicase activity (measured by optical tweezers). Below we describe the steps to making these measurements, the analysis of the data, and interpretation of the results.

5.1 Protein Expression and Fluorescent Labeling We prepare UvrD labeled with a FRET donor–acceptor pair at positions on the protein shown to exhibit a large change in FRET efficiency upon transition from “closed” to “open” state (Fig. 1) (Jia et al., 2011; Lee & Yang, 2006). We use standard maleimide chemistry for fluorescence labeling (Joo & Ha, 2008), where the cysteines (Cys) in the protein react specifically to the maleimide form of the dyes AlexaFluor555 (donor) and AlexaFluor647 (acceptor) (Molecular Probes, Eugene, OR). A UvrDΔCys(A100C,A473C) mutant was constructed with all six native cysteine residues replaced with serine and with two separate mutations at position 100 and 473 substituting alanine to cysteine. The protein was labeled stochastically with a 1:1 ratio of donor and acceptor dyes. Details on protein expression and purification are reported elsewhere (Jia et al., 2011).

5.2 DNA Hairpin Construct We utilize a hairpin assay (Dumont et al., 2006) to monitor helicase unwinding at high resolution. As shown in Fig. 1, a molecule containing a DNA hairpin is tethered between two optically trapped beads. To monitor helicase-catalyzed unwinding, the assay is performed at a constant force below that required to unfold the hairpin mechanically and the extension of the tethered molecule is monitored with the instrument in force clamp. For every 1 bp unwound by the helicase, 2 nt are released and extend the tethered molecule, mechanically amplifying the unwinding signal (Dumont et al., 2006). For example, at a force of 10 pN, each bp unwound generates a 0.8-nm extension change signal. The DNA hairpin construct consists of three DNA fragments that are ligated together (Fig. 7): a “hairpin” (HP) flanked by a “right handle”

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Fig. 7 Construction of DNA hairpin substrate. (A) Schematic depicting the major steps involved in preparing the DNA construct. (B) A representative force–extension curve of the final construct (black), showing models of the folded (magenta) and unfolded (blue) states.

(RH) and a “left handle” (LH) that serve as functionalized linkers that attach to the trapped beads. RH is made from a 1.5-kb PCR-amplified section of the pBR322 plasmid (New England Biolabs) using a 50 digoxigenin-modified forward primer and a reverse primer containing one abasic site and a long 50 overhang (Table 1). The digoxigenin moiety is used to link this end of the construct to an antidigoxigenin-coated bead. The overhang consists of a 19-nt poly-dT loading site for helicase binding immediately adjacent to the abasic site, followed by 29 nt that anneal to a complementary sequence in HP. LH is synthesized from a different PCRamplified section of pBR322 using a 50 -biotin-modified primer. The biotin moiety allows this end of the construct to bind to a streptavidin-coated bead. HP is a single long oligonucleotide containing the complementary sequence to the LH overhang on its 50 end, followed by the complementary sequence to the RH overhang and a 153-nt self-complementary sequence (Table 1). When self-annealed and ligated to LH and RH, HP makes an 89-bp hairpin stem capped by a (dT)4 tetraloop.

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Table 1 Oligonucleotides for Constructing Hairpin Oligonucleotide Sequence (IDT Format, 50 to 30 )

LH forward primer

/5Biosg/ TGA AGT GGT GGC CTA ACT ACG

LH reverse primer

CAA GCC TAT GCC TAC AGC AT

RH forward primer

/5Phos/TTG AAA TAC CGA CCG CTC AGC TAT CAG CCT TTT TTT TTT TTT TTT TTT /idSp/CTC TGA CAC ATG CAG CTC CC

RH reverse primer

/5DigN/ CAA CAA CGT TGC GCA AAC T

Hairpin insert

/5Phos/CCT GGG GCT GAT AGC TGA GCG GTC GGT ATT TCA AAA GTC AAC GTA CTG ATC ACG CTG GAT CCT AGA GTC AAC GTA CTG ATC ACG CTG GAT CCT ATT TTT AGG ATC CAG CGT GAT CAG TAC GTT GAC TCT AGG ATC CAG CGT GAT CAG TAC GTT GAC TT

5.2.1 Synthesis and Purification of LH and RH 1. For PCR synthesis of LH, mix 35 μL of nuclease-free water, 5 μL of forward primer (10 μM concentration), 5 μL of reverse primer (10 μM), 2 μL of pBR322 template DNA (10 ng/μL) (NEB), 3 μL DMSO, and 50 μL 2 Phusion HF Master Mix (NEB) for a final volume of 100 μL. 2. For PCR synthesis of RH, mix 35 μL of nuclease-free water, 5 μL of forward primer (10 μM concentration), 5 μL of reverse primer (10 μM), 2 μL of pBR322 template DNA (10 ng/μL), 3 μL DMSO, and 50 μL 2 Phusion HF Master Mix for a final volume of 100 μL. 3. Run PCR on both reaction mixes. We use the following program: (1) 98°C for 30 s, (2) 98°C for 10 s, (3) 59°C for 10 s, (4) 72°C for 33 s, (5) repeat steps 2–4 30, (6) 72°C for 5 min, and (7) 4°C forever. 4. Purify PCR products following the QIAquick PCR purification kit “spin protocol” (Qiagen). Add 30 μL of elution buffer instead of 50 μL for a more concentrated solution. These PCR products can be verified by gel electrophoresis. 5.2.2 Digestion of LH and Removal of the 50 -Phosphate 5. Add the following to 30 μL of LH: 2 μL of PspGI restriction enzyme (20 units total; NEB) and 3.5 μL of CutSmart 10  buffer (NEB).

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6. Incubate the LH reaction mix at 75°C for 1 h. 7. Add the following to the 35 μL of digested LH: 4 μL 10 Antarctic phosphatase buffer (NEB), 1 μL Antarctic phosphatase (5 units; NEB). 8. Incubate the new LH reaction mix at 37°C for 30 min, then 80°C for 2 min to inactivate the phosphatase. 9. Purify the digested LH using the QIAquick PCR purification kit as before. 5.2.3 Ligation of LH and RH to HP 10. Measure the concentrations of LH and RH using a Nanodrop UV–vis spectrophotometer (Thermo Scientific), which requires only 1 μL of each solution. 11. Mix the three DNA components in an equimolar ratio (1:1:1) to a final volume of 32 μL. 12. Add 4 μL of 10  T4 DNA ligase buffer (NEB) and 4 μL of T4 DNA ligase (1600 units; NEB) for a final volume of 40 μL. 13. Ligate at RT (22°C) for 1 h and then heat to 65°C for 15 min to inactivate T4 ligase. 14. Run the final product on a 1% agarose gel with no ethidium bromide for 70 min. 15. Incubate the gel in a solution of 50 mL 0.5  TBE and 15 μL 10,000 GelGreen (Biotium) for  20 min. 16. Image the gel using Dark Reader Transilluminator (Clare Chemical Research) and cut out the appropriate band (3.4 kb) using a clean razor blade. Place this gel slice in a previously weighed 1.5-mL tube. 17. Purify the final ligated construct from the agarose gel slices using QIAEX II gel extraction kit (Qiagen). Add 30 μL of elution buffer instead of 50 μL for a more concentrated solution.

5.3 Sample Flow Chamber During measurements, we use custom laminar flow sample chambers to assemble protein–DNA complexes in situ. Chambers consist of a central channel with two adjacent laminar fluid streams that do not mix (Qi et al., 2013) (Fig. 8). In the experiments described below (Fig. 9), the lower stream contains ATP (10 μM) but no protein, while the upper stream contains UvrD (10 nM) but no ATP. By moving the sample chamber stage, we can move the optical traps from one stream to the other in a few seconds (typically 2 s). This allows us to assemble the protein–DNA complex first

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Fig. 8 Laminar flow cell layout. (A) Chambers are assembled from a piece of parafilm melted between two #1 coverslips. The top coverslip has inlet holes cut into it. The parafilm has patterned into it two outer channels for flowing in beads, and a central channel consisting of two laminar flow streams for assembling complexes in situ. Two glass capillaries connect the top and bottom channels to the central channel and are used to dispense beads. (B) An assembled flow chamber is mounted on a custom aluminum bracket into which tubing mates with the inlet holes. The tubing assembly consists of PE tubing inserted into short lengths of Tygon tubing threaded through set screws screwed into the bracket. (C) Photograph of a flow chamber.

(in the upper stream), then initiate unwinding (in the lower stream) in a controlled fashion. The laminar flow chambers are made by melting a piece of parafilm between two #1 microscope cover glasses (Fig. 8; Fisher Scientific) as described previously (Qi et al., 2013). Briefly, we use a CO2 laser engraver (VSL2.25; Universal Laser Systems) to cut four inlet and outlet holes into one cover glass. We use the same laser engraver to cut three channels into the parafilm. The top and bottom channels are used to flow in antidigoxigenin- and streptavidin-coated beads, respectively. A small flow of beads enter into the central channel via glass capillaries (ID ¼ 0.0250 mm, OD ¼ 0.10 mm; King Precision Glass) that are embedded in the melted parafilm and shunt the top and bottom channels to the central channel. The central channel consists of two separate streams from two central inlets, which merge smoothly and maintain a sharp interface due to the laminar, nonturbulent flow.

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Fig. 9 Simultaneous measurement of UvrD conformation and activity. (A) Steps for in situ nucleoprotein complex assembly. The flow chamber consists of three channels. The top (yellow) and bottom (green) channels contain antidigoxigenin (ADig) and DNAcoated streptavidin (DNA) beads, respectively, and the central measurement channel consists of two parallel laminar flow streams containing 10 nM protein (blue) and 10 μM ATP (red). During an experiment, we carry out the following steps in sequence: (1) trap an ADig bead dispensed out of the top capillary, (2) move to the bottom capillary and trap a DNA bead, (3) calibrate the traps, (4) form a tether by bringing the beads into contact and pulling apart (and optionally taking a F–x curve), (5) move to the protein stream and load UvrD by incubating for 15 s, and (6) move to the ATP stream, turning on the fluorescence excitation. (B) Model of UvrD conformational switching, in which the closed (open) state corresponds to translocation into (away from) the DNA fork, on opposite strands of the hairpin. (C) Simultaneous measurement of donor (green, top panel) and acceptor (red) fluorescence intensity, FRET efficiency (middle panel), and hairpin base pairs unwound (bottom panel) during UvrD unwinding. A correlation is seen between the “closed,” high FRET state and hairpin unwinding (grayshaded time intervals) and “open,” low FRET state and hairpin rezipping (unshaded time intervals). Figure adapted from Comstock, M. J., Whitley, K. D., Jia, H., Sokoloski, J., Lohman, T. M., Ha, T., & Chemla, Y. R. (2015). Direct observation of structure-function relationship in a nucleic acid-processing enzyme. Science, 348(6232), 352–354 with permission from AAAS.

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In order to prevent proteins from adsorbing to the cover glasses, we passivate the glass surfaces with polyethylene glycol (PEG) in situ using the following protocol modified from Ha et al. (2002): 1. Clean and sonicate cover glasses in 3 M KOH for 20 min prior to assembling the flow cell. 2. Assemble the flow cell. 3. Flow a solution of 1% (v/v) N-(2-aminoethyl)-3-aminopropyltrimethoxysilane (United Chemical Technologies, Bristol, PA) and 5% (v/v) acetic acid in methanol through all channels and let incubate for 15 min. 4. Rinse the channels with methanol and water, flow a solution of 25% (w/v) methoxy-PEG-succinimidyl valerate (MW 5000; Laysan Bio, Arab, AL) in 0.1 M sodium bicarbonate, and let incubate for 4 h. 5. Rinse the flow cell with copious amounts of Type 1 deionized water and dry with nitrogen gas. Flow chambers are stored dry in the dark at 4°C. The sample chamber is mounted on a custom bracket with screw-in adapters (Fig. 8) for flowing in buffers and samples via polyethylene (PE) tubing (427406, BD Intramedic). During measurements, we use motorized syringe pumps (PHD Ultra Nanomite 703601 and PHD Ultra Remote Infuse/ Withdraw Programmable 703107, Harvard Apparatus) to inject buffers and samples loaded in glass syringes (PTFE Luer Lock, 81320, Hamilton) into the chamber. Each inlet is controlled by a separate pump. We use a programmable syringe pump for the central channel for better control over the laminar flow. Typical volumetric flow rates are 100 μL/h, corresponding to a linear flow rate of 140 μm/s.

5.4 Sample and Buffer Preparation for Optical Trap–smFRET Assay Here we describe the sample preparation steps for our example experiment. First, we incubate the DNA hairpin construct with streptavidin-coated polystyrene microspheres (810-nm diameter, SVP-08-10, Spherotech) in a 1:1 DNA:bead ratio for 1 h to allow biotin–streptavidin linkages to form. Second, we prepare antidigoxigenin antibody-coated polystyrene beads as follows: 1. Add 40 μL of 1% w/v Protein G microspheres (880-nm diameter, PGP08-5, Spherotech) into 160 μL 1  PBS buffer + 0.01% Tween 20. Tween helps prevent bead aggregation. 2. Wash the beads twice by centrifuging at 6000 rcf for 1 min, remove the supernatant, and resuspend with PBS + Tween solution.

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3. Add 10 μL of 1 mg/mL antidigoxigenin antibody (11333089001, Roche) into the bead solution after the last resuspension. 4. Rotate or shake the solution for 30–60 min. 5. Wash the beads 2  as in step 2 to remove excess antibody. Store in final wash (PBS + Tween). Measurements are conducted in 35 mM Tris (pH 8.0), 20 mM NaCl, 5 mM MgCl2, and 2% glycerol. We use an oxygen scavenging system to increase the lifetimes of the fluorophores and the tethers (Ha, 2001; Landry, McCall, Qi, & Chemla, 2008)—1.2% glucose, 1 mg/mL glucose oxidase (SigmaAldrich, St. Louis, MO), and 0.13 mg/mL catalase (EMD Millipore, Billerica, MA)—and a triplet-state quencher to prevent fluorophore blinking (Rasnik, McKinney, & Ha, 2006)—1 mg/mL Trolox (Sigma-Aldrich, St. Louis, MO).

5.5 Optical Trap–smFRET Measurement The first step for this experiment is to tether the DNA hairpin between two trapped beads. We first trap an antidigoxigenin bead flowing out of the top capillary (Fig. 9A), then translate the sample stage such that the traps are near the bottom capillary and capture a streptavidin–DNA bead in the second trap. The traps are then calibrated by the standard method (BergSørensen & Flyvbjerg, 2004; Neuman & Block, 2004), measuring the beads’ Brownian motion to extract the conversion factors αx,y between bead position and QPD output (in nm/V) and the trap stiffnesses κx,y (in pN/nm). The QPD outputs a zero-force offset voltage that we find depends on trap position; we map this offset voltage, moving the trapped beads together after calibration. We then form a tether by bringing the beads into contact, waiting a short time, and moving them apart. The detection of a force (>4 pN) indicates a tether has been formed. We often take a force– extension curve to ensure that a single molecule is tethering the beads and to examine the quality of the DNA construct (Fig. 7B). The tethered DNA molecule is held at a constant force (5–13 pN) as it is moved into the channel containing UvrD to load protein onto the DNA construct. After incubating in the UvrD channel for 15 s, the tether is then moved back into the channel containing ATP, and the excitation laser is turned ON. Trap and fluorescence data are acquired. Fig. 9C shows a time trace obtained from such an experiment. The raw QPD bead position data are collected at the interlacing rate (66 kHz) and boxcar averaged to 133 Hz. As the helicase unwinds the DNA hairpin,

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the released ssDNA extends the tethered molecule. Under a constant force, this “slack” is taken up by moving the traps further apart. To determine the number of base pairs that are unwound, we first convert the change in the AOM1 RF drive frequency (in MHz) into a trap displacement (in nm). This conversion factor is obtained a priori by taking images of trapped beads at different RF frequencies with the visible system CCD camera, and determining the positions of the bead centers. The conversion factor for our setup is 123 nm/MHz, but will be different depending on the details of the setup. Next, we convert the trap displacement (in nm) to base pairs (bp) unwound by dividing by the extension of the 2 released nucleotides per bp unwound at the force of the measurement. We use the worm-like chain model, with interphosphate distance 0.59 nm/nt, persistence length 1 nm, and stretch modulus 1000 pN, which fits well to our force–extension curve of the mechanically unzipped DNA hairpin (Fig. 7B). The simultaneous fluorescence signal (Fig. 9C) is analyzed as follows. We integrate the photon counts from the APDs over a set time window (typically 10 ms) and determine photon emission rates (in kHz; top panel) by dividing the integrated counts by the time window. We determine the FRET efficiency E (middle panel) from the standard formula: E ¼ IA =ðIA + ID Þ, where IA and ID are the fluorescence intensities (photon rates) of the acceptor and donor dyes, respectively (Ha, 2001). We subtract the background fluorescence signals obtained after both dyes have photobleached to obtain IA and ID. Fig. 9C illustrates the power of using multiple simultaneous methods to measure biomolecular dynamics. The data trace of FRET efficiency reveals that UvrD does not remain in one conformational state during its activity but rather switches between open and closed states (compare shaded and unshaded regions). Meanwhile, the trap data (bottom panel) show not only periods of unwinding of the hairpin (shaded regions), but also periods during which the duplex rezips (unshaded regions). The simultaneous measurement of conformational state and UvrD activity demonstrates a clear correlation between the two types of dynamics. In the closed (high FRET) state the helicase is on average unzipping DNA, while in the open (low FRET) state the DNA is rezipped. This correlation provides new insights into the functional roles of the open and closed states of UvrD. Based on prior structural and singlemolecule work, we believe the two conformational states correspond to UvrD translocating on opposing strands of the hairpin (Comstock et al., 2015). In the closed state, the helicase translocates 30 –50 into the DNA fork,

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unwinding it, while in the open state it moves 30 –50 on the opposing strand, away from the fork, allowing the duplex to rezip in UvrD’s wake (Fig. 9B).

ACKNOWLEDGMENTS We thank members of the Chemla and Comstock laboratories for scientific discussion. Funding was provided by NSF grants MCB-0952442 (CAREER to Y.R.C.), PHY1430124 (Center for the Physics of Living Cells to Y.R.C.), RC-105094 (to M.J.C.), and NIH grant R21 RR025341 and R01 GM120353 (to Y.R.C.).

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Moffitt, J. R., Chemla, Y. R., Aathavan, K., Grimes, S., Jardine, P. J., Anderson, D. L., & Bustamante, C. (2009). Intersubunit coordination in a homomeric ring ATPase. Nature, 457(7228), 446–450. Moffitt, J. R., Chemla, Y. R., Izhaky, D., & Bustamante, C. (2006). Differential detection of dual traps improves the spatial resolution of optical tweezers. Proceedings of the National Academy of Sciences of the United States of America, 103(24), 9006–9011. Neuman, K. C., & Block, S. M. (2004). Optical trapping. The Review of Scientific Instruments, 75(9), 2787–2809. Qi, Z., Pugh, R. A., Spies, M., & Chemla, Y. R. (2013). Sequence-dependent base pair stepping dynamics in XPD helicase unwinding. eLife, 2, 1–23. Rasnik, I., McKinney, S. A., & Ha, T. (2006). Nonblinking and long-lasting single-molecule fluorescence imaging. Nature Methods, 3(11), 891–893. Ritchie, D. B., & Woodside, M. T. (2015). Probing the structural dynamics of proteins and nucleic acids with optical tweezers. Current Opinion in Structural Biology, 34, 43–51. Sirinakis, G., Ren, Y., Gao, Y., Xi, Z., & Zhang, Y. (2012). Combined versatile high-resolution optical tweezers and single-molecule fluorescence microscopy. Review of Scientific Instruments, 83(9), 093708. Stryer, L., & Haugland, R. P. (1967). Energy transfer: A spectroscopic ruler. Proceedings of the National Academy of Sciences of the United States of America, 58(2), 719–726. Suksombat, S., Khafizov, R., Kozlov, A. G., Lohman, T. M., & Chemla, Y. R. (2015). Structural dynamics of E. coli single-stranded DNA binding protein reveal DNA wrapping and unwrapping pathways. eLife, 4, 1–23. van Dijk, M. A., Kapitein, L. C., van Mameren, J., Schmidt, C. F., & Peterman, E. J. G. (2004). Combining optical trapping and single-molecule fluorescence spectroscopy: Enhanced photobleaching of fluorophores. Journal of Physical Chemistry B, 108, 6479–6484. van Mameren, J., Modesti, M., Kanaar, R., Wyman, C., Wuite, G. J. L., & Peterman, E. J. G. (2006). Dissecting elastic heterogeneity along DNA molecules coated partly with Rad51 using concurrent fluorescence microscopy and optical tweezers. Biophysical Journal, 91(8), L78–L80. van Mameren, J., Wuite, G. J. L., & Heller, I. (2011). Introduction to optical tweezers: Background, system designs, and commercial solutions. Methods in Molecular Biology, 783, 1–20. Veigel, C., & Schmidt, C. F. (2011). Moving into the cell: Single-molecule studies of molecular motors in complex environments. Nature Reviews. Molecular Cell Biology, 12(3), 163–176. Visscher, K., Brakenhoff, G. J., & Krol, J. J. (1993). Micromanipulation by multiple optical traps created by a single fast scanning trap integrated with the bilateral confocal scanning laser microscope. Cytometry, 14, 105–114. Visscher, K., Gross, S. P., & Block, S. M. (1996). Construction of multiple-beam optical traps with nanometer-resolution position sensing. Journal of Selected Topics in Quantum Electronics, 2(4), 1066–1076. Whitley, K. D., Comstock, M. J., & Chemla, Y. R. (2017). High-resolution “fleezers”: Dual-trap optical tweezers combined with single-molecule fluorescence detection. Methods in Molecular Biology, 1486, 183–256.

CHAPTER SEVEN

Integrating Optical Tweezers, DNA Tightropes, and Single-Molecule Fluorescence Imaging: Pitfalls and Traps J. Wang*, J.T. Barnett*, M.R. Pollard†, N.M. Kad*,1 *School of Biosciences, University of Kent, Canterbury, Kent, United Kingdom † DFM A/S, Kongens Lyngby, Denmark 1 Corresponding author: e-mail address: [email protected]

Contents 1. Introduction 2. Elongating Bundled DNA for Imaging 2.1 DNA Combing 2.2 DNA Tightropes 3. Integrating Laser Tweezers Into Biological Experiments 3.1 Introduction 3.2 Trapping With Nanoprobes 4. Controlling and Detecting the Nanoprobe 4.1 Using an AOD to Create Three Traps 4.2 Detection Strategies 5. Applying the Nanoprobe to Biological Study Systems 5.1 Measuring the Tension of a DNA Tightrope 5.2 Measuring Interactions With Single Proteins 6. Conclusions and Outlook References

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Abstract Fluorescence imaging is one of the cornerstone techniques for understanding how single molecules search for their targets on DNA. By tagging individual proteins, it is possible to track their position with high accuracy. However, to understand how proteins search for targets, it is necessary to elongate the DNA to avoid protein localization ambiguities. Such structures known as “DNA tightropes” are tremendously powerful for imaging target location; however, they lack information about how force and load affect protein behavior. The use of optically trapped microstructures offers the means to apply and measure force effects. Here we describe a system that we recently developed to enable individual proteins to be directly manipulated on DNA tightropes. Proteins bound to DNA can be conjugated with Qdot fluorophores for visualization and also Methods in Enzymology, Volume 582 ISSN 0076-6879 http://dx.doi.org/10.1016/bs.mie.2016.08.003

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directly manipulated by an optically trapped, manufactured microstructure. Together this offers a new approach to understanding the physical environment of molecules, and the combination with DNA tightropes presents opportunities to study complex biological phenomena.

1. INTRODUCTION Proteins and substrates exist in a physical milieu where they are subject to processes such as Brownian motion, diffusion, and external force. The contribution of these factors toward the overall mechanism requires the properties of the individual components to be determined for extrapolation to the ensemble system. One major biological process that requires understanding at this level is diffusion to target, and particularly so for targets on DNA. Based on diffusion models, it is possible to predict the association time of a protein to its target site (Berg, 1993). However, Lac repressor was found to locate its target site faster than predicted by a three-dimensional (3D) encounter model (Riggs, Suzuki, & Bourgeois, 1970). Facilitated diffusion was postulated to explain this discrepancy and involves a change in the mechanism of target search from being wholly 3D to also include onedimensional (1D) diffusion along the backbone of the DNA (Adam & Delbr€ uck, 1968; Winter, Berg, & von Hippel, 1981). For many years, numerous investigators set out to test this hypothesis (Halford & Marko, 2004) prior to the application of relatively new single-molecule methods. These methods permit the observer to track a single molecule labeled with a fluorophore as it moves toward its target site. Early investigations proved the existence of 1D sliding (Blainey, van Oijen, Banerjee, Verdine, & Xie, 2006; Tafvizi et al., 2008), hopping (Bonnet et al., 2008) which is longdistance jumps in position along a single DNA strand and also a combination of all of these mechanisms (Hughes et al., 2013; Kad, Wang, Kennedy, Warshaw, & van Houten, 2010). Also clear was that some proteins were capable of directional motor movement (van Oijen et al., 2003). To study such systems at the single-molecule level, first it requires DNA to be elongated. In Fig. 1 we show an image of a field of λ-DNA molecules stained with YOYO-1 dye. λ-DNA (the genome of bacteriophage lambda) consists of 48,502 base pairs; with a base pair separation of 0.34 nm, this equates to a contour length of 16.5 μm. It is apparent in Fig. 1 that the molecules in the image have collapsed into a bundled conformation. The radius  pffiffiffiffiffiffiffiffi of a bundle can be calculated using random flight theory r ¼ l  K ,

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Fig. 1 Image of YOYO-1 stained bacteriophage λ-DNA molecules. This DNA has a contour length of 16 μm; however, in solution, the DNA collapses into bundles. These diffuse through the field of view and are also seen to change shape.

where l is the contour length and K is the Kuhn length (twice the persistence length), for λ-DNA this is 1.3 μm. Comparing this to the point spread function of a typical fluorophore (Anderson, Georgiou, Morrison, Stevenson, & Cherry, 1992; Thompson, Larson, & Webb, 2002), it is apparent that measuring motion within this bundle is extremely challenging. Furthermore, the bundle will move, so the positional accuracy is adversely affected. Here we describe how we construct DNA tightropes to achieve elongated DNA structures and how this can be combined with optically trapped probes for more sophisticated experiments.

2. ELONGATING BUNDLED DNA FOR IMAGING 2.1 DNA Combing DNA can be laid onto an activated surface (Allemand, Bensimon, Jullien, Bensimon, & Croquette, 1997), usually polystyrene or polymethylacrylate by a method known as combing (Bensimon et al., 1994). This is achieved by either retracting an activated slide through a solution of DNA or flowing DNA across a surface (Crut et al., 2005; Deen et al., 2015; Lyon, Fang, Haskins, & Nie, 1998). Typically performed at low pH to achieve better binding, the DNA can then be examined using total internal reflection fluorescence microscopy. This reduces the amount of background fluorescence permitting fluorescently labeled proteins on DNA to be clearly imaged. In addition to this approach the DNA can be attached by one end and flow applied to elongate the DNA such as used to study the DNA repair protein

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human oxoguanine DNA glycosylase 1 (Blainey et al., 2006) or DNA replication (Lee et al., 2006). Although more complex, this approach reduces, but does not eliminate, interactions between DNA-bound proteins and the surface. Visually, however, it is unclear whether the proteins are attached to the DNA or surface because both are in the same focal plane.

2.2 DNA Tightropes This method exploits the simplicity of combing but raises the DNA from the surface without the need for flow. Suspending individual DNA molecules between surface-immobilized beads ensures that anything visible within the focal plane above the surface must be bound to DNA and not the surface. To facilitate imaging, it is necessary to optimize illumination such that only the tightropes are excited and the rest of the solution receives minimal light. This is achieved by using oblique angle fluorescence excitation microscopy (Hughes et al., 2013; Konopka & Bednarek, 2008; Tokunaga, Imamoto, & Sakata-Sogawa, 2008). To create DNA tightropes we use the following protocol: 1. Plasma clean 22  40 mm coverslips (Harrick PDC-32G, Ithaca, NY) for 2 min per slide. 2. Build a flow chamber as described in Fig. 2. 3. Block the chamber overnight by flowing in mPEG5000 (Sigma Aldrich) pH 8.2, wash with water, and then block overnight again with 50 mM Tris–HCl, 10 mg/mL BSA, 0.1% Tween20. 4. Flow in 5 μm silica beads (precoated overnight with 350 μg/mL poly-Llysine (Sigma Aldrich)), until bead separation is on average 20 μm. 5. Wash flow chamber with 200 μL buffer and check bead density again to ensure that beads have not been washed from the surface. In that event recoat beads with poly-L-lysine before adding to flow chamber (see Fig. 2 legend for details on constructing a flow chamber; also see Kad et al., 2010). 6. Attach a syringe pump (AL-1000 World precision instruments) to one flow chamber outlet and to the other a microcentrifuge tube using polyethylene tubing (PE90; BD Medical, Oxford, UK). 7. Backflow the buffer from the syringe into the microcentrifuge tube to ensure an airtight connection. 8. Flow in 100 μL of 15 nM λ-DNA, this will bring the DNA into the flow chamber. Apply forward and backward flow to run the DNA across the surface beads for at least 20 min.

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Fig. 2 Design of flow cells for creating tightropes. To create flow chambers a plasma cleaned glass coverslip (#1.5) is attached to a glass microscope slide using an in-house cut double-sided adhesive gasket (UK industrial tapes) of depth 180 μm. The glass slide is predrilled (using a diamond bur; Precision Dental, London, UK) to allow the attachment of small diameter tubing and sealed using UV curable adhesive (NOA68, Thorlabs, Ely, UK) after insertion. The gasket design is shown in the lower panel, indicating the positions of ports used for tightropes and for nanoprobe injection. Image taken with permission from Simons, M., Pollard, M. R., Hughes, C. D., Ward, A. D., van Houten, B., Towrie, M., et al. (2015). Directly interrogating single quantum dot labelled UvrA2 molecules on DNA tightropes using an optically trapped nanoprobe. Scientific Reports, 5, 18486.

9. Flush the flow chamber with 200 μL of imaging buffer and if required 100 μL of 1 nM YOYO-1 dye to visualize the DNA. During experiments, it is usual to exclude the addition of YOYO-1 dye since it is known that anything in focus at the height of the beads will be bound to DNA.

3. INTEGRATING LASER TWEEZERS INTO BIOLOGICAL EXPERIMENTS 3.1 Introduction Optical trapping, also known as laser tweezing, permits physical manipulation of single molecules. This approach requires a high-energy laser beam focused into a fluid sample to interact with particles of higher refractive index than their surrounding solution. As a consequence the particles, typically polystyrene or silica beads, become fixed near the focal point of the laser

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beam. Deviation from the focal point results in a restoring force that is proportional to the distance from the center of the trap position over a region of approximately 500 nm. This Hookean response results in a very simple linear correlation between displacement and force. Therefore, not only are laser tweezers useful for physical placement of molecules but also their response to force. The application of laser tweezers for studying molecular motors was initially established with kinesin (Svoboda, Schmidt, Schnapp, & Block, 1993) and myosin (Finer, Simmons, & Spudich, 1994; Guilford et al., 1997; Molloy, Burns, Kendrick-Jones, Tregear, & White, 1995). The latter experiments elegantly suspend individual actin filaments between optically trapped beads, pull them taut (Dupuis, Guilford, Wu, & Warshaw, 1997), and then use this platform to investigate the properties of single myosin molecules attached to a solid surface (Fig. 3). These experiments have revolutionized biology by permitting the load response of single molecules to be determined (Kad, Patlak, Fagnant, Trybus, & Warshaw, 2007; Kad, Trybus, & Warshaw, 2008; Veigel, Molloy, Schmitz, & Kendrick-Jones,

Fig. 3 The three-bead assay. This assay, used primarily for studies of the molecular motor myosin, involves plating a low density of myosin onto a coverslip surface that is precoated with 3-μm silica beads. 1-μm silica beads are trapped and used to capture the ends of an actin filament, which is then tautened. When the actin is brought into contact with the surface-bound myosin and the position of the trapped beads detected, a clear drop in variance is seen. This is due to the additional contribution of the stage to the stiffness of the system. In addition, myosin will move the actin which is seen as a deflection from the baseline of the bead position.

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2003; Veigel, Schmitz, Wang, & Sellers, 2005). Such measurements are not possible at the bulk level and also permit the contribution of the single molecule in the context of the ensemble to be determined. Such scaling measurements are integral to the construction of nanobiotechnological devices (Hess, 2011) and for the understanding of complex biological systems. Beyond molecular motors the study of DNA-based enzymes has also been facilitated by the use of laser tweezers. This has typically taken the form of suspending single DNA molecules between beads in much the same way as actin has been used for studying myosin (Hilario, Amitani, Baskin, & Kowalczykowski, 2009; Qi, Pugh, Spies, & Chemla, 2013; Skinner, Baumann, Quinn, Molloy, & Hoggett, 2004; Wang, Yin, Landick, Gelles, & Block, 1997). However, recently more elaborate applications of laser tweezers have been used to study the localization and affinity of proteins for DNA. These include looping one DNA molecule around another (Noom, van den Broek, van Mameren, & Wuite, 2007; van Loenhout et al., 2013) or incorporating atomic force microscope probes (Huisstede, Subramaniam, & Bennink, 2007; Shon & Cohen, 2016).

3.2 Trapping With Nanoprobes 3.2.1 Fabrication and Preparation of Nanoprobes Optical trapping has traditionally involved the use of spherical beads that are isotropically suspended in the traps. This means that the movement of the beads is limited to simple translation and that the surface available for interacting with molecules is large and cannot be easily controlled. To overcome these limitations, we and others have used structured particles that can be optically trapped. As shown in Fig. 4 we use a triangular structure with three

Fig. 4 The structure of the nanoprobe. Fabricated using EBL from SU8 the nanoprobe is a triangular structure with three cylinders located at each vertex. Three laser traps are used to position the probe each focused on a different cylinder. The probe is a long thin blade structure, and the nanoprobe is visible by fluorescence as well as bright-field microscopy. Image taken with permission from Simons, M., Pollard, M. R., Hughes, C. D., Ward, A. D., van Houten, B., Towrie, M., Botchway, S. W., Parker, A. W., & Kad, N. M. (2015). Directly interrogating single quantum dot labelled UvrA2 molecules on DNA tightropes using an optically trapped nanoprobe. Scientific Reports, 5, 18486.

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trapping points and a protruding tip (Pollard et al., 2010). This permits accurate 3D positioning of the protrusion such that molecules can be manipulated with greater precision. The “nanoprobe” structures were fabricated using electron beam lithography as follows. To etch the nanoprobes a chrome layer followed by an SU8 photoresist layer is applied to a silicon wafer, baked, and then exposed (Pollard et al., 2010). This process yielded 4000 nanoprobes per segment on a silicon wafer. Correct release of the nanoprobes from the wafer is pivotal to enable subsequent experiments to be performed efficiently. First, the chrome layer is dissolved using ceric ammonium nitrate (CAN) pipetted directly onto the nanoprobe containing segment. The duration of exposure to etchant is controlled in order to leave some chrome connecting the nanoprobes to the wafer. This permits subsequent washing steps without loss of nanoprobes; any remaining CAN residue in the wafer region containing nanoprobes is gently washed away using high-purity water several times. Residual CAN was found to inhibit downstream steps, and washing also removes any free particulates that can contaminate the flow chamber and interfere with trapping experiments. After several washes the nanoprobes can be released with a more vigorous rinse with buffer containing BSA and Tween20 (Simons et al., 2015). This step prevents aggregation of the nanoprobes which are then collected and kept for use in the flow cell. 3.2.2 Flow Chamber Adaptations for Improved Experimental Yield For introduction of the nanoprobes into the flow chamber, it is necessary to avoid contact with the walls of the tubing, which was found to bind avidly to the nanoprobes. Therefore to add nanoprobes, a separate hole was drilled into the flow chamber requiring a specific design with a separate channel for nanoprobes (Fig. 2). This design not only avoids the use of polyethylene tubing during addition of nanoprobes but also creates a channel for storage of nanoprobes within the flow chamber. Nanoprobes are injected using a glass Hamilton syringe, and the channel can be sealed simply using commercially available sticky tape.

4. CONTROLLING AND DETECTING THE NANOPROBE Laser tweezers are relatively simple to construct, requiring only the back aperture of a high numerical aperture (>1.2) objective lens to be slightly overfilled with a stable laser. To obtain the correct beam size requires

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the use of one or more telescopes in the optical path. Controlling the position of the laser tweezers can be achieved by a number of schemes; however, they all share a common point that the turning mechanism is conjugated with the back focal point of the objective lens. This ensures a stable wide region for trapping, for a much more extensive discussion on the tweezer setup used (see Sung, Sivaramakrishnan, Dunn, & Spudich, 2010). Later we discuss the control mechanism used for our system (Simons et al., 2015) and the detection scheme.

4.1 Using an AOD to Create Three Traps In order to manipulate the laser-trapping beam laterally across the imaging plane, acoustic optical deflectors (DTD-274HD6M, IntraAction) are inserted into the laser beam path to deflect the beams with varying acoustic frequency (Molloy, 1998). In such a way, three optical traps can be created to coincide spatially with the vertex positions of the nanoprobe. These are generated by rapidly (40 kHz) repositioning the laser between each position significantly faster than the nanoprobe response frequency of 30 Hz (Pollard et al., 2010). Removal of the IR-blocking filter permits an image of the trap position to be projected onto the camera, although this should only be performed when necessary to prevent damage to the camera. Once a floating nanoprobe is found in solution, adjust the orientation of the traps to match it, and note their position. Turn off the IR-trapping laser. Move the trap position marker over the nanoprobe and adjust the focus to be slightly above the nanoprobe. Turning on the IR laser should capture the nanoprobe. Since the nanoprobes are located in the side channel of the flow chamber (Fig. 2), they will need to be navigated to an appropriate DNA tightrope. This is a delicate operation requiring movements over millimeters, and touching anything in the flow chamber will likely result in the nanoprobe sticking. At high IR laser powers, it is possible to guide the nanoprobe over the surface beads. However, to image the tightropes simultaneously, the trapping and imaging planes need to be offset in the z-axis. This is achieved by moving the focusing lens for the optical tweezers. Once a tightrope is found, the lens is returned to its correct position bringing the fluorescence and trapping planes into parfocality. At this point, it is necessary to align the detection apparatus with the center of the nanoprobe vertices and to perform some calibrations of the trap stiffness. For the latter, position detection is required.

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4.2 Detection Strategies Classical detection of trapped objects requires either a position sensitive detector (PSD) or a quadrant photodiode (QPD). These record the position of the trapped object at the back focal plane of the microscope condenser. The recorded signal is an interference signal between the scattered light from the bead and incident trapping beam. This detection approach relies on the linear relationship between the center of mass of the interference pattern and the relative position between the trapping beam and the trapped bead; however, this scheme is not suitable for tracking multiple beads simultaneously. The output of a PSD, usually a voltage signal, will be affected by parameters such as power of the beam and its shape and size; therefore, careful calibration is required. We adopt a more flexible detection method of bead tracking by using image-based position detection which allows us to track multiple beads by following their movement. Previous studies (Belloni, Monneret, Monduc, & Scordia, 2008; Keen, Leach, Gibson, & Padgett, 2007; Otto, Gutsche, Kremer, & Keyser, 2008) have shown that image-based tracking can provide similar performance compared with PSD and QPD detection schemes with the added advantage of multiple trap detection with flexible controls. Multiple moving objects can be tracked using real-time updated region of interests (ROIs) across the image. With PSD or QPD detectors offering up to hundreds of kHz, image-based trapping cannot compete in terms of detection bandwidth. However, for most position measurements, a fast camera with a carefully positioned ROI will suffice. 4.2.1 Optics for Image-Based Detection The position detection path and the laser-trapping beam are built around a wide-field microscope. As shown in Fig. 5, a high-power white LED light source is placed above the sample stage of the microscope in a transmission configuration to provide bright-field illumination. A Hamamatsu ORCA-Flash 2.8 CMOS camera (Hamamatsu Photonics, Hamamatsu, Japan) with 1920  1440 pixels is used as the image sensor to acquire brightfield microscopic images. The imaging signal collected by the microscope objective, OBJ, is focused by a doublet achromatic lens (L1, f ¼ 200 mm) to form a conjugate imaging plane where a mechanical pupil (A) is placed to control the aperture size. A pair of achromatic lenses L2 (f ¼ 100 mm) and L3 (f ¼ 50) form a 4F configuration to relay the image at A to the image sensor. The trapping laser is a 5W 1070-nm ytterbium fiber-coupled laser that is separated from the detection path by a dichroic beam splitter. The laser beam is focused by the objective to the sample plane, SP, within the

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Fig. 5 Layout of the camera-based position detection scheme. CMOS, Hamamatsu’s Scientific CMOS Camera, ORCA-Flash 2.8; DBS, dichroic beam splitter; A, aperture; L1, L2, and L3, achromatic lens with focal length of 200, 100, and 50 mm, respectively; LED, brightfield light source; NBF, notch-blocking filter; OBJ, microscope objective; SP, sample plane; the dashed arrows indicate the positions of image plane and image conjugate planes.

field of view of the camera. A notch-blocking filter is used to stop the back reflection of the trapping beam reaching the camera. In this way, the camera can deliver the image of the SP with the trapping laser focus in the middle. Each pixel on the camera covers an area of 120  120 nm2 in the SP; therefore, an ROI of 16  16 pixels produces a detection window of 1.92  1.92 μm2. With this ROI, the ORCA-Flash 2.8 camera runs at a maximum frame rate of 1 kHz. The pixel size is calibrated by using a graticule (Graticules Ltd., Tonbridge, UK). To initially locate objects for trapping the full field of view is used. ROIs are then adopted to isolate individual objects floating in the solution. 4.2.2 Centroid Detection We describe here the use of 1.5-μm diameter silica beads as a trapped object; however, the procedure for a nanoprobe vertex is identical. When the bead is trapped by the laser, the image of the bead appears bright in the middle, providing an image with high contrast and small size. The center of mass is calculated by using the centroid approach in LabView™. N X N  X

Cx ¼

i  I ij



i¼1 j¼1 N X N   X I ij i¼1 j¼1

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The centroid position in x (Cx) takes the sum of the product of the intensity of each pixel in a column (Iij) and that column position in x (i). This is repeated for every column in the ROI (i ¼ 1 to N). Finally the sum of all of these values is expressed relative to total intensity of the image. This calculation is repeated for the y-axis but for each row. We normally threshold the image to offset problems of low contrast and to remove any distractions from background features in the flow chamber. Using this detection system clear steps (20 kb) DNA substrates with lesions and other extrahelical structures inserted at defined positions. DNA derived from bacteriophage λ (λ-DNA) is a high quality long (48.5 kb) DNA substrate that is frequently used in singlemolecule studies. Here we provide detailed protocols for site-specific incorporation of recombinant sequences and extrahelical structures into λ-DNA. We also describe Methods in Enzymology, Volume 582 ISSN 0076-6879 http://dx.doi.org/10.1016/bs.mie.2016.08.006

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2017 Elsevier Inc. All rights reserved.

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how to assemble DNA curtains, and how to collect and analyze single-molecule observations of lesion recognition by MMR proteins diffusing on these DNA curtains. These protocols will facilitate future single-molecule studies of DNA transcription, replication, and repair.

1. INTRODUCTION The highly conserved DNA mismatch repair (MMR) system recognizes and repairs nucleotide misincorporation events and extrahelical lesions that are introduced during DNA replication and homologous recombination. In eukaryotes, repair is initiated by one of two heterodimeric MutS homolog (Msh) complexes, Msh2–Msh6 and Msh2–Msh3. Both complexes form sliding clamps on DNA and scan the genome for a partially overlapping but distinct spectrum of DNA lesions (Erie & Weninger, 2014; Jiricny, 2013; Kantelinen et al., 2010; Lee et al., 2014; Li, 2008). Msh2–Msh6 primarily recognizes single-nucleotide mismatches and small insertion– deletion loops (IDLs) (Jiricny, 2013; Reyes, Schmidt, Kolodner, & Hombauer, 2015). Msh2–Msh3 also recognizes some single-nucleotide mismatches as well as IDLs involving one or more unpaired nucleotides (Harrington & Kolodner, 2007; Srivatsan, Bowen, & Kolodner, 2014; Surtees & Alani, 2006). After the lesion is recognized, MutL homolog (Mlh) complexes—Mlh1–Pms1 (in yeast) or Mlh1–Pms2 (in humans)— are recruited to the repair site. Mlh complexes also form sliding clamps on homoduplex DNA and harbor a mismatch-dependent endonuclease activity that is critical for MMR (Kadyrov, Dzantiev, Constantin, & Modrich, 2006; Kadyrov et al., 2009, 2007). Following Mlh recruitment to the vicinity of the lesion, exonuclease 1 (Exo1) excises a long tract of DNA from the lesion containing strand. Finally, the gapped DNA is filled in by DNA polymerases (Kunkel & Erie, 2005). MMR requires a spatially and temporally controlled assembly of repair enzymes at the lesion. Single-molecule imaging and manipulation have proven especially amenable for deciphering how these dynamic assemblies recognize and excise the lesion (Erie & Weninger, 2014; Lee et al., 2014). For example, single-molecule approaches have been used to describe how Msh proteins scan both naked and chromatinized DNA for lesions, how Mlh proteins are recruited to the Msh–lesion complex, and how Exo1 excises the lesion (Brown et al., 2016; Gorman et al., 2007; Jeon et al., 2016; Lee et al., 2014; Myler et al., 2016; Qiu et al., 2012). These studies

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Fig. 1 A strategy for inserting extrahelical structures into λ-DNA. Step 1: A nicking cassette is inserted into the λ-phage genome in vivo. Step 2: Recombinant λ-DNA is purified. Step 3: Extrahelical structures are introduced via a nicking enzyme-based oligonucleotide insertion strategy. B and D represent incorporated biotinylated and digoxigeninlabeled oligonucleotides, respectively. Step 4: The resulting DNA substrates are assembled into microfluidic DNA curtains and imaged via single-molecule microscopy.

frequently require site-specific incorporation of mismatches or other lesions on a long (>20 kb) DNA substrate. DNA isolated from bacteriophage λ (λ-DNA) is a convenient source of high quality long (48.5 kb) DNA. However, introducing specific DNA sequences, tertiary structures, and chemical modifications into λ-DNA remains technically challenging. In this chapter, we provide detailed protocols for rapidly modifying and purifying recombinant λ-DNA for single-molecule imaging (Fig. 1). We use in vivo recombineering to target site-specific segments of the λ-phage genome with >90% efficiency, abrogating the need for restriction motifs and ligation. A nicking enzyme (nickase)-based strategy is used to incorporate mismatches and other structures at defined positions along the DNA substrate. Furthermore, we describe how to use these substrates for assembling DNA curtains, a high-throughput single-molecule fluorescence imaging approach (Gallardo et al., 2015). We anticipate that these protocols will be broadly useful for both ensemble and single-molecule studies that require site-specific modification of long DNA substrates.

2. MATERIALS 2.1 Buffers 1. L1 buffer: 300 mM NaCl; 100 mM Tris–HCl [pH 7.5]; 10 mM EDTA; 0.2 mg mL
1 bovine serum albumin (BSA; fraction V, Sigma-Aldrich). 2. L2 buffer: 30% polyethylene glycol (PEG 6000) (w/v); 3 M NaCl. 3. L3 buffer: 100 mM NaCl; 100 mM Tris–HCl [pH 7.5]; 25 mM EDTA. 4. L4 buffer: 4% sodium dodecyl sulfate (SDS) (w/v). 5. L5 buffer: 3 M potassium acetate [pH 5.5]. 6. QBT buffer: 750 mM NaCl; 50 mM MOPS [pH 7.0]; 15% isopropanol (v/v); 0.15% Triton X-100 (v/v).

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7. QC buffer: 1.0 M NaCl, 50 mM MOPS [pH 7.0]; 15% isopropanol (v/v). 8. QF buffer: 1.25 M NaCl, 50 mM Tris–HCl [pH 8.5]; 15% isopropanol (v/v). 9. TE buffer: 10 mM Tris–HCl [pH 8.0]; 1 mM EDTA. 10. SM buffer: 50 mM Tris–HCl [pH 7.5]; 100 mM NaCl; 8 mM MgSO4. 11. 10  alkaline agarose gel electrophoresis buffer: 500 mM NaOH; 10 mM EDTA. 12. Neutralization buffer: 1 M Tris–HCl [pH 7.5]; 1.5 M NaCl. 13. Lipids buffer: 10 mM Tris–HCl [pH 8.0]; 100 mM NaCl. Filter through a 0.22 μm syringe filter (Olympus) and store at room temperature. 14. BSA buffer: 40 mM Tris–HCl [pH 7.8]; 1 mM DTT; 1 mM MgCl2; 0.2 mg mL
1 BSA. Filter through a 0.22 μm syringe filter and use the same day as experiment. 15. Imaging buffer: BSA buffer with 50 mM NaCl; 1–5 nM YOYOI (Life Tech); 500 units of catalase (Sigma-Aldrich); 70 units of glucose oxidase (Sigma-Aldrich); and 1% glucose (w/v). 16. PBS buffer: 10 mM phosphate [pH 7.2]; 138 mM NaCl; 2.7 mM KCl. Autoclave and stored at room temperature.

2.2 Recombineering 1. 2. 3. 4.

Micropulser (Bio-Rad). Micropulser electroporation cuvettes (Bio-Rad). Lab shaking incubator (New Brunswick). Optima XE-90 Ultracentrifuge (Beckman Coulter).

2.3 Assembling DNA Curtains 1. TE2000 Eclipse Inverted Microscope (Nikon) modified for prism TIRF illumination, with a 488 nm laser (Coherent), a Nikon Plan Apo 60 objective (Nikon), a 638 dichroic beam-splitter (Chroma), a 500 long-pass filter (Chroma), and two EM-CCD cameras (Andor iXon DU897) (Finkelstein & Greene, 2011; Gallardo et al., 2015). 2. Quartz Dove Prism (Tower Optical). 3. Index matching immersion oil (type FF, Cargile). 4. Syringe pump (KD Scientific). 5. Rheodyne HPLC 6 port switching valve (Kinesis). 6. Microfabricated quartz slides made via UV lithography techniques (Gallardo et al., 2015).

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7. λ-DNA (New England Biolabs). 8. Liposomes for surface functionalization and passivation. Liposomes are made by combining three different lipid solutions (purchased as powder from Avanti Polar Lipids) and diluting them in chloroform. DOPC (100 mg mL
1) (1,2-dioleoyl-sn-glycero-3-phosphocholine) (#850375P), DOPE-mPEG2k (100 mg mL
1) (1,2-dioleoyl-snglycero-3-phosphoethanolamine-N-[methoxy(polyethylene glycol)2000]) (#880130P), and DOPE-biotin (25 mg mL
1) (1,2dioleoyl-sn-glycero-3-phosphoethanolamine-N-(cap biotinyl)) (#870273P). Liposome stock solutions are made by mixing a molar ratio of 97.7% DOPC, 2.0% DOPE-mPEG2k, and 0.3% DOPE-biotin (Finkelstein & Greene, 2011). 220 μL of liposome mixture is then evaporated overnight in a vacuum desiccator, and rehydrated in 2 mL lipids buffer for 4 h. The liposomes are sonicated using a microtip sonicator (Qsonica Q700) at 15% power for three 90 s intervals. This ensures small, unilamellar vesicles (see Note 1). Sonicated products are filtered through a 0.22 μm syringe filter and stored at 4°C. 9. Streptavidin (Life Technologies) stored as 1 mg mL
1 solution in H2O at
20°C. 10. Digoxigenin monoclonal antibody (Life Technologies). 11. Goat antirabbit IgG polyclonal antibody (Immunology Consultants Laboratory #GGHL-15A). 12. PFA tubing 0.0200 ID (IDEX). 13. Flowcell injection connectors constructed from various IDEX fittings. NanoPort 10–32 coned ports, FingerTight III 10–32 fitting, 1/1600 replacement ferrule, hex short 1/1600 fitting. 14. Disposable Luer-lock connector syringes (BD Scientific).

2.4 Imaging MMR Proteins 1. 2. 3. 4. 5.

YOYO1 dye: stored as 1 mM stock in DMSO at
20°C (Invitrogen). Glucose oxidase type II from Aspergillus niger (Sigma-Aldrich). Catalase from bovine liver (Sigma-Aldrich). SiteClick Qdot 705 antibody labeling kit (ThermoFisher). Affinity purified polyclonal rabbit anti-HA (Immunology Consultants Laboratory #RHGT-45A-Z) conjugated to quantum dots (QDs). Label antibodies using the SiteClick Qdot 705 antibody labeling kit (Life Technologies #S10454) per the manufacturers directions with slight modification (see Note 2). Store conjugates at 4°C for up to 3 months.

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3. METHODS We developed an improved nickase-based strategy for inserting oligonucleotides at precise positions along λ-DNA (Fig. 1). First, a cassette that includes three consecutive BspQI recognition motifs separated by 20-bp spacers is cloned into the λ-phage genome. To facilitate this process, we designed a series of cassettes that target two nonessential regions within the phage genome (Fig. 2). Each nickase cassette is flanked by 200-bp of homology to λ-DNA. λ-Red-based recombineering is used to construct recombinant λ-DNA rapidly with >90% efficiency (Sharan, Thomason, Kuznetsov, & Court, 2009). Recombinant lysogens are identified via colony PCR and through growth sensitivity at a restrictive temperature (42–45°C). Recombinant phage is induced by a heat shock and the λ-DNA is purified from packaged phage particles. Below, we provide detailed protocols for each of these steps.

3.1 Recombineering into λ-Phage 3.1.1 Preparing the Nickase Cassette A linear, double-stranded DNA PCR product that contains a nicking cassette can be incorporated at one of three defined positions along the length of λ-DNA (defined as position A, B, or C). The nickase cassette contains a triple Nt.BspQI nickase recognition motif and an antibiotic resistance gene flanked on either end by 200 bps of sequence homology to λ-DNA. The identity of the flanking λ-DNA homology arms determines which of the three positions (A, B, or C) will be targeted for nickase cassette

Fig. 2 Incorporating recombinant DNA into the λ-phage genome. (a) A schematic representation of the 48.5 kb-long λ-DNA. Nonessential regions of the phage genome are indicated with a dotted line. We routinely insert recombinant DNA in one or more of the three positions marked A, B, and C. (b) Colony PCR is used to screen recombinant lysogens. For each position, the smaller bands indicate a successful insert. (c) Accurate recombineering is confirmed via an NcoI digest of the purified λ-DNA substrates.

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incorporation. We have designed a set of helper plasmids that can be used as PCR templates to amplify the desired nicking cassette with the specified λ-DNA homology arms. These plasmids (pIF251, pIF253, and pIF254) are available upon request. 1. Amplify the nickase cassette by PCR from the helper plasmid of choice (pIF251, pIF253, and pIF254 for positions A, B, and C, respectively) using Taq DNA polymerase and the specified primers (Tables 1 and 2). 2. Degrade the template by adding 1 U of restriction enzyme DpnI (NEB #R0176s) and incubating at 37°C for 1 h. Table 1 Nicking Cassette Preparation Replacement Position λ-Position (Relative to cosL)

Plasmid Template

Primers Forward

Reverse

A

21,251–22,750 bp

pIF251

AD027

AD028

B

33,498–34,685 bp

pIF253

AD012

AD013

C

46,446–47,862 bp

pIF254

AD016

AD017

Each nicking cassette is generated by PCR using the indicated template and primers. Templates are available on request.

Table 2 Nickase and Primer Site Sequences Name Sequence

IF003

/5Phos/AGG TCG CGG CC/3Bio

IF004

/5Phos/GGG CGG CGA CCT/3Dig

AD012

AGT CTG GAT AGC CAT AAG TG

AD013

GTA ACC ACA TAC TTC CTG CC

AD016

GCA GTC TGT CAG TCA GTG CG

AD017

CGA GGG CAT TGC AGT AAT TG

AD027

GCT ACC ACC ATG ACT AAC GC

AD028

GGA TAT CAG AGC TAT GGC TC

AD006

TTTTTTTTTTTTTTTTTTTTTTTTTTTTTT TCTTCCCTTGGTGCGATCGCTCTTCG

3  Nt.BspQI nickase motif

GCT CTT CAT GCA TGC GGC CGC TCT TCC CAT GGT GCG ATC GCT CTT CGG

Sequences are read 50 –30 . IF003 and IF004 were HPLC purified. AD006 was PAGE purified.

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3. Purify the PCR product by gel extraction and resuspend in Milli-Q H2O to a final concentration of 100–150 μg μL
1. Removing residual plasmid DNA is essential, as even trace amounts can cause false positives during recombineering. 3.1.2 Electroporation and Recombineering 1. Grow a 5 mL LB culture of λ-phage lysogen strain IF189 + pKD78 overnight at 30°C in the presence of 10 μg μL
1 chloramphenicol. pKD78 is an arabinose-inducible helper plasmid that encodes the λ-red recombineering proteins (Datsenko & Wanner, 2000). IF189 was created by infecting infect Escherichia coli LE392MP cells with λ-DNA (λc1857 Sam7) using MaxPlax λ-phage packaging extracts (Epicenter). 2. The following day, use 350 μL of cells to inoculate a fresh 35 mL culture of LB supplemented with 10 μg mL
1 chloramphenicol. 3. When the cells reach an OD600  0.5, induce the λ-red recombinase system by adding 2% L-arabinose (w/v) (GoldBio #20-108) and incubate for an additional 1 h at 30°C. 4. Harvest the cells by centrifugation at 4500 RCF for 7 min. 5. To make the cells electrocompetent, wash 3  in ice-cold Milli-Q H2O followed by centrifugation and resuspension in 200 μL of H2O after each wash. Keep the electrocompetent cells on ice and use them immediately for the recombineering reaction (see Note 6). 6. Combine 50–150 ng of the nickase cassette (prepared above) with 50 μL of electrocompetent cells and transfer them into ice-cold 0.1 cm cuvettes. Electroporate the PCR product into the cells at 18 kV cm
1. 7. Immediately resuspended the cells in 1 mL of SOC and then transfer to culture tubes containing 10 mL LB broth. 8. After a 4 h outgrowth at 30°C, plate 100 μL of the culture onto LB agar plates containing a low concentration of the appropriate antibiotic, and incubate overnight at 30°C. 9. The following morning, pick colonies and check for successful incorporation of recombinant DNA via colony PCR (Fig. 2).

3.2 Purifying λ-DNA The following procedure describes the overexpression and harvesting of recombinant phage capsids, and subsequent purification of approximately 1 mL of 200–500 ng μL
1 of λ-DNA. Lytic λ-phage lysogen growth is induced at 45°C. This temperature denatures the temperature-sensitive λ-repressor (cI857ind 1), which initiates phage amplification and packaging.

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An additional amber mutation in the S gene (Sam 7) delays cell lysis and produces large burst sizes (>200 phage capsids per cell), ultimately maximizing the yield of recombinant λ-DNA. 3.2.1 Induction of Lytic Growth 1. Grow a colony of recombinant λ-phage lysogen in 50 mL of LB broth with the appropriate antibiotic overnight at 30°C. 2. Use 10 mL of this starter culture to inoculate 500 mL of LB the following morning. When the flask reaches an OD600  0.6, rapidly raise the temperature to 42°C by swirling in a preheated water bath. 3. Once the temperature reaches 42°C, induce lytic growth by transferring the culture to a 45°C shaking incubator for 15 min. 4. After 15 min, lower the incubator temperature to 37°C and continue shaking for 2 h. 3.2.2 Purification 1. Harvest the cells by centrifugation at 3000 RCF for 30 min and decant the supernatant (see Note 8). 2. Resuspend the cell pellet in 10 mL of SM buffer. 3. Add 2% chloroform (v/v) and lyse the cells by incubating at 37°C for 30 min while shaking at 200 rpm. 4. Degrade residual bacterial genomic DNA and RNA by adding 50 ng μL
1 DNaseI (Sigma #D2821) and 30 ng μL
1 RNaseA (Sigma #R6513) to the lysate and incubate for an additional hour at 37°C. 5. Centrifuge the lysate for 15 min at 6000 RCF and at 4°C. Collect the supernatant and dilute with 40 mL of SM buffer. This clarified lysate contains soluble phage capsids. 6. Add 10 mL ice-cold L2 buffer and precipitate the phage capsids by incubating on ice for 1 h. 7. Harvest the phage capsids by centrifugation at 10,000 RCF for 10 min at 4°C. Decant the supernatant. 8. Resuspend the pellet with 3 mL of L3 buffer and 3 mL of L4 buffer. 9. Phage capsid proteins are digested by incubating with 100 ng μL
1 of proteinase K (NEB #P8012S) for 1 h at 55°C, which liberates the λ-DNA. 10. Precipitate the SDS with 3 mL of L5 buffer and clarify the resulting cloudy solution by centrifugation at 15,000 RCF for 30 min at 4°C. Collect the supernatant.

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11. Preequilibrate a Qiagen tip-500 column (Qiagen #10262) with 30 mL QBT buffer. 12. Titrate the solution to pH 7.0 using 1 M MOPS [pH 8.0]. Pass the soluble λ-DNA over the column. 13. Wash the column with 30 mL QC buffer and elute with 15 mL of QF buffer. 14. Precipitate the λ-DNA by adding 10.5 mL of 100% isopropanol. DNA will precipitate as a long stringy web. 15. Collect the precipitate with a bent Pasteur pipette or pipette tip. 16. Rinse the spooled pellet in 70% ethanol and dissolve in TE buffer to a final DNA concentration of 200–500 ng μL
1. 17. An NcoI restriction digest can be used to confirm the quality of the recombinant λ-DNA (Fig. 2c).

3.3 Inserting Oligonucleotides into DNA Substrates Recombinant λ-DNA is nicked with Nt.BspQI (nickase) enzyme, which creates closely spaced breaks in the top strand of the nickase cassette (Fig. 3a). Alternatively, Nb.BspQI can be used to modify the bottom strand. Short single-stranded DNA fragments are melted out and replaced by a synthetic oligonucleotide by annealing slowly and ligating overnight. Excess oligonucleotides and enzymes are then removed by gel filtration. Restriction enzyme motifs (NcoI, NotI) that are within the replacement positions are used to verify proper insertion of the synthetic oligonucleotide (Fig. 3b). Complete ligation of both strands can be further confirmed with alkaline agarose electrophoresis (Fig. 3c). 1. In a 250 μL reaction, incubate 25 μg of recombinant λ-DNA with 50 U of Nt.BspQI (NEB #R0644s) and 1  buffer 3.1 (NEB #B7203) at 55°C for 1 h. Add 1 U of proteinase K (NEB #P8107S) for 1 h, at 55°C to halt the reaction, and increase the temperature to 70°C for 20 min. 2. Add a 500-fold molar excess of the desired insert oligonucleotide, along with a 250-fold excess of cosL and cosR-complimentary oligonucleotides (IF003 and IF004; Table 2). The cosL and cosR oligonucleotides are modified with biotin or digoxigenin for assembling DNA curtains (Finkelstein & Greene, 2011; Gallardo et al., 2015). 3. Slowly reduce the temperature from 70 to 22°C in a thermocycler at a rate of
0.5°C min
1. 4. Add 1000 U of T4 DNA ligase and 1 mM ATP. Incubate overnight at room temperature. 5. Remove 50 μL of the final reaction for insertion verification. Save the rest for gel filtration.

Fig. 3 Constructing λ-DNA with an internal single-stranded DNA flap. (a) Schematic of the nickase-based oligonucleotide replacement reaction. B and D represent biotinylated and digoxigenin-labeled oligonucleotides, respectively. (b) A restriction digest can be used to quantify oligonucleotide replacement rapidly. Inserting a 50 -ssDNA flap, but not a homoduplex oligonucleotide, produces a 2.7-kb fragment. Homoduplex and mock-treated λ-DNA are further digested into 2 and 0.7-kb fragments (0.7-kb band not shown). (c) A denaturing (alkaline) agarose gel confirms insertion and religation of the λ-DNA substrates. Note that the top and bottom DNA strands are separated only for the 50 -ssDNA flap substrate.

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6. Inactivate 250 μL of the above reaction with a final concentration of 1 M NaCl. Load the solution onto a 120 mL Sephacryl S-1000 column (GE #17-0476-01) in TE running buffer plus 150 mM NaCl to separate the modified λ-DNA from excess oligonucleotides and enzymes. 3.3.1 Diagnostics The nicking cassette contains NcoI and NotI motifs for measuring oligonucleotide incorporation efficiency. If possible, design the insertion reaction such that successful incorporation of the oligonucleotide into the λ-DNA abolishes either restriction enzyme motif. Under these conditions, incorporation of the insert can be verified by performing a restriction enzyme digest. The restriction fragments are resolved on a native 0.8% agarose gel. Additional analysis on a denaturing alkaline agarose gel can be used to verify proper ligation of the modified λ-DNA (Fig. 3). Nicking with Nt.BspQI produces a distinct ladder of bands on an alkaline agarose gel (Fig. 3c). Ligation seals the nicks that cause these bands. 3.3.2 Alkaline Agarose Gel Diagnostics 1. Take 5 μL of the sample set aside for insertion verification and run on a 0.6% alkaline agarose gel made with 1 alkaline agarose gel electrophoresis buffer. Run the gel at 20 V for 20 h at 4°C. 2. Rock the gel in neutralization buffer for 45 min at room temperature. 3. Stain by soaking in 10 mg mL
1 ethidium bromide dissolved in 1  alkaline agarose gel electrophoresis buffer for 30 min at room temperature. 4. Destain the gel by soaking in H2O for 30 min at room temperature. 5. Visualize DNA using a Typhoon FLA 9500 laser scanner (GE) (Fig. 3).

3.4 Assembling DNA Curtains 1. Assemble flowcells and prepare connectors as previously described (Finkelstein & Greene, 2011). A diagram of the UV lithography generated features as well as double-tethered DNA curtains can be seen in Fig. 1 (Gallardo et al., 2015). Briefly, a single DNA curtain barrier is comprised of a deposited line of Chromium (Cr) separated 13 μm away from circular Cr pedestals. These features are generated on a quartz flowcell via UV lithography. λ-DNA is tethered between these features via biotin/streptavidin and digoxigenin/α-digoxigenin antibody interactions, respectively. A lipid bilayer is deposited on the surface of the flowcell to both facilitate the attachment of λ-DNA to the bilayer and to passivate the surface of the flowcell.

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2. Wash the flowcell with H2O. Equilibrate the flowcell with lipids buffer (see Note 13). 3. Dilute 40 μL of liposome solution into 960 μL of lipids buffer. Inject diluted liposomes into the flowcell in three rounds with 10 min of incubation between each round. 4. Wash the flowcell in lipids buffer and incubate for at least 30 min for lipid healing. This will create a uniform lipid bilayer for surface passivation and λ-DNA attachment/organization. 5. Dilute 15 μL of goat antirabbit polyclonal antibody (Immunology Consultants Laboratory #GGHL-15A) into 285 μL of lipids buffer and inject into the flowcell. Incubate for 10 min. The antibody will attach nonspecifically to the chromium features including the pedestals. 6. Wash the flowcell in BSA buffer and immediately inject 2.5 μL of digoxigenin monoclonal antibody (Life Technologies #700772) diluted into 250 μL BSA buffer. Incubate for 10 min. The primary antibody will bind to the secondary antibody located on the pedestals. This will capture the digoxigenin functionalized end of the λ-DNA, allowing for tethering at both ends of the DNA molecule. 7. Dilute 30 μL of streptavidin into 270 μL of BSA buffer (0.1 mg mL
1) and inject into the flowcell. Incubate for 10 min. Streptavidin will bind the biotinylated lipids and serve as a mobile tether for the functionalized λ-DNA. 8. Wash the flowcell with BSA buffer. Dilute 200 μL of functionalized λ-DNA into 800 μL of BSA buffer. Inject λ-DNA into the flowcell in three rounds with 5 min of incubation between each round. 9. Connect the flowcell to a syringe containing 10 mL of imaging buffer loaded onto a syringe pump (Fig. 4). Flow imaging buffer through the flowcell at a rate of 100 μL min
1. This flow rate provides enough force to extend the λ-DNA to be captured at the pedestal. 10. Attach the flowcell to the microscope stage using stage clips. Set the prism onto the flowcell with a drop of index matching oil. The polished faces of the prism should be in line with the excitation laser. The flow cell is ready for imaging (Fig. 4).

3.5 Imaging DNA MMR Proteins on DNA Curtains 1. Acquire data through the use of Nikon Elements (Nikon) or other software. Microscope settings for acquisition are 10 MHz camera readout mode, 300 EM gain, 5  conversion gain, and 200 ms frame rate.

Fig. 4 Visualizing Msh2–Msh3 binding to an extrahelical ssDNA flap. (A) Cartoon representation (left) and picture (right) of a microscopemounted flowcell. The picture highlights the microfluidic connectors and the quartz prism. (B) Distribution of Msh2–Msh3 molecules on flapcontaining λ-DNA. The black line is a Gaussian fit to the data (n ¼ 503). The center of the peak corresponds to the expected location of the lesion (20 kb from the top DNA barrier, error is SD). Inset: Seven representative λ-DNA molecules (light gray vertical lines) with flap-bound Msh2–Msh3 (black points). Msh2–Msh3 recognizes lesions via (C) 1D diffusion along the DNA or (D) a direct encounter (3D collision). Each panel shows a cartoon illustration (top), kymograph (middle), and single-particle trajectory of Msh2–Msh3 (black) binding a DNA flap (30 -ssDNA flap; marked as 30 ). Asterisk indicates transient binding by a second Msh2–Msh3. In both cases, Msh2–Msh3 releases the flap and continues to diffuse on homoduplex λ-DNA.

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2. If included in the imaging buffer, image YOYOI with 20 mW laser power (see Note 15). Use YOYOI to focus on the surface as well as to fine-tune the TIRF angle. 3. The Msh2 subunit encodes an HA epitope tag between amino acids 644 and 645. Label Msh2–Msh3 with anti-HA QDs (Brown et al., 2016; Gorman et al., 2007). Briefly, coincubate QDs and protein at a 1:1 molar ratio (150 nM protein and QDs) in BSA buffer for 15 min on ice followed by dilution to a final concentration of 5–10 nM in imaging buffer. 4. Inject the diluted protein–QD mixture into the flowcell using a 6-port switching valve and incubate with DNA curtains for 5–10 min. 5. After incubation, flush excess QDs and all non-DNA-bound proteins using buffer flow. 6. Terminate buffer flow and begin data acquisition. 7. Visualize QD-labeled Msh2–Msh3 with 20 mW laser power. QDs can be imaged indefinitely without photobleaching. Export raw data TIFFs without compression and save for analysis.

3.6 Analyzing Single-Molecule Traces 1. Analyze the raw data using ImageJ (NIH), or another image processing software. Track fluorescent particles to subpixel resolution by fitting the fluorescent intensity of a single QD to a two-dimensional Gaussian. An objective comparison of several particle-tracking strategies is available online (Chenouard et al., 2014). Our homemade ImageJ script is also available on request. 2. Generate digital trajectories and binding histograms using tracking information (Fig. 4). Trajectories and histograms can be created using Matlab (Mathworks) or another computing environment. 3. Calculate the 1D mean squared displacements (MSD) of individual molecules using:
n X 1 N ðyi + n
yi Þ2 MSDðnΔt Þ ¼ N
n i¼1 where N is the total number of frames in the trajectory, n is the number of frames for a given time interval, Δt is the time between frames (time interval), and yi is the molecule position at frame i. 4. Calculate the diffusion coefficient of individual molecules by fitting a line through a plot of the MSDs for the first 10 time intervals using the following formula: MSDðΔt Þ ¼ 2DΔt where D is the apparent 1D diffusion coefficient.

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4. NOTES 1. Sonicated liposomes can be stored for up to 2 weeks at 4°C. Liposomes will continually condense in solution over the storage period. After approximately 2 weeks, the liposomes stop producing uniform DNA curtains. 2. After conjugating antibodies to QDs, purify the labeled QDs from the free antibody with a Sephacryl S-300 column (GE Life Sciences) or a similar gel filtration column in PBS buffer. 3. The targeting inserts can be prepared in advance and stored at
20°C. 4. Although recombineering can be performed with a minimum of 40 ng μL
1 targeting insert λ-DNA, we saw much higher recombineering efficiency when the DNA was >100 ng μL
1. 5. An OD600 0.5 is optimal for arabinose induction of the λ-red recombinase system encoded on pKD78. The recombineering efficiency will markedly decrease at an OD600 > 0.6. 6. Make electrocompetent cells fresh and use them as soon as they have been prepared. Add SOC immediately after electroporation, and transfer cells to LB and the incubator as soon as possible. 7. Although it is often enough to allow cells to grow out for 4 h after electroporation, it is highly recommended to allow the cells to grow for 24 h. This greatly increases the number of colonies per plate and increases the likelihood of successfully recombineered lysogens. 8. The λ-phage cell pellet can be flash frozen here and stored at
80°C for up to 1 week. 9. It is critical not to raise the temperature above 45°C during the heat shock of λ-phage lysogens. Monitor the LB temperature with a thermometer, as higher temperatures may affect the yield of purified λ-DNA. 10. The phage capsid pellet is faint white and slightly opaque; it can be very difficult to see on the walls of a 50 mL conical tube. Mark the tube position within the rotor when harvesting the phage capsid as to not obstruct the pellet from view. 11. Purification of >500 ng μL1 λ-DNA from these modified λ-phage lysogens may be expected but a minimum concentration of 200 ng μL
1 should be achieved before attempting the nicking reaction. 12. When diagnosing the insertion efficiency, include the proper mock (no insert) and homoduplex insert controls.

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13. When injecting solutions into the flowcell, be careful not to introduce bubbles, as they will destroy the lipid bilayer. To mediate this problem, make drop to drop connections with all syringes and push bubbles out of the flowcell by injection from the opposite port. 14. When aligning the TIRF angle, decrease the laser power to 3 h. To facilitate injection of PEG solution, cut a 1-cm piece of flexible, narrow Tygon tubing to use as a seal between the pipette tip used to handle the PEG solution and the hole in the glass chamber assembly. 39. Wash out the unbound PEG with 500 μL of TN buffer.

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40. Store for up to 3 weeks at 4°C in a humid chamber (i.e., a Parafilmsealed Petri dish containing a sheet of sterile tissue and a small amount of Milli-Q water). 5.1.7 Final Chamber Preparation 41. Cut the bottom off of two microfuge tubes (0.625 mL eppendorf tube for instance) and glue over the through-hole in the glass using UV-cured glue and a UV-curing lamp. 5.1.8 Functionalization of Biotinylated PEG Prior to use of a surface, the biotin groups at the end of the PEG chains must be functionalized with streptavidin. 1. Prepare a stock solution of 2 mg/mL streptavidin in TN buffer. This solution can be stored at 4°C for 3 months or divided into 10 μL aliquots and stored at 20°C. 2. Take an aliquot of the above solution and dilute 10-fold using TN buffer. 3. Introduce into flow cell and incubate 10 min at room temperature. 4. Rinse with 500 μL of TN buffer.

6. ANTIDIGOXIGENIN-DERIVATIZED POLYSTYRENECOATED GLASS SURFACES 6.1 Materials Spin coater (MTI Corporation, Richmond, CA, USA, VTC-100) Hydroxymethyldisilazane (HMDS) solution (Microchem, Westborough, MA, USA, MCC Primer 80/20) Toluene (AnalaR NORMAPUR grade, VWR, 28676.297) Polystyrene (Sigma, St. Louis, MO, USA, 331651) Phosphate-buffered saline (PBS), pH 7.4 Antidigoxigenin (Roche, Basel, Switzerland, 11 333 089 001) BSA (Roche, 10 775 835 001) Blocking buffer (40 mM KHEPES, pH 8, 100 mM NaCl, 8 mM MgCl2, 0.1% (w/v) Tween-20, 10 mM β-mercaptoethanol) Temperature block A second convenient approach for preparing treated surfaces is provided, which is based on a hydrophobic polystyrene layer. These surfaces can readily be coated with antidigoxigenin and used, in conjunction with standard streptavidin-coated magnetic beads, to tether the same DNA constructs as those used for fluorescence experiments. These surfaces reliably provide for a very high density of DNA tethers with minimal surface interactions

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and are thus convenient for working out biochemical aspects of experiments of interest. Unfortunately, fluorescent species present in polystyrene typically precludes use of these surfaces in fluorescence assays. The protocol presented below is identical to that presented in SLIDE CLEANING up to and including step 22. Then, 23. Spin-coat onto each slide a layer of HMDS. Place slide in spin coater, cover slide with an uninterrupted layer of HMDS solution, spin 5 s at 500 rpm, and then spin 30 s at 4000 rpm. 24. Anneal 10 min on a temperature block heated to 110°C. 25. Spin-coat onto each slide a layer of polystyrene (0.1%, w/v in toluene). Place slide in spin coater, cover slide with an uninterrupted layer of toluene solution, spin 5 s at 500 rpm, and then spin 30 s at 4000 rpm. 26. Assemble into chambers as described in Section 5.1.7. 6.1.1 Functionlization of Polystyrene-Coated Glass Coverslip 27. Resuspend 200 μg of antidigoxigenin in 2 mL of 1 PBS; make 20 μL aliquots, and store at 20°C. 28. Dilute an aliquot of antidigoxigenin 10-fold with PBS buffer, fill flow cell with solution, and incubate overnight at 37°C. 29. The next morning, wash out unbound antidigoxigenin with 1 mL of 1 PBS. 30. Inject into flow cell blocking buffer supplemented with 10 mg/mL BSA and incubate 1½–2 h at 37°C. Longer incubation may cause BSA to precipitate. 31. Replace solution with blocking buffer containing 0.1 mg/mL BSA. Store for up to 1 month at 4°C in a humid chamber (i.e., a parafilmsealed Petri dish containing a sheet of sterile tissue and a small amount of Milli-Q water).

7. PREPARATION OF DNA 7.1 Materials DNA plasmid Competent DH5α bacteria (e.g., from MAX Efficiency cells, ThermoFisher Scientific, 18258012) Luria broth (LB) Ampicilin (Sigma, A9393) DNA Maxiprep kit (Macherey-Nagel, D€ uren, Germany, NucleoBond Xtra Maxi Plus)

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DNA PCR and Gel Cleanup kit (Macherey-Nagel, NucleoSpin Gel & PCR Cleanup) Restriction enzymes (New England Biolabs) T4 DNA ligase 400,000 U/mL (New England Biolabs, Ipswich, MA, USA, M0202S) Expand HiFidelity PCR System (Roche, 11 732 650 001) Digoxigenin-11-dUTP, 1 mM (Roche, 11 093 088 910) Biotin-16-dUTP, 1 mM (Roche, 11 093 070 910) dNTP mix, 10 mM each (Roche, 11 814 362 001) Standard wet lab equipment (PCR machine, tabletop centrifuge, agarose gel electrophoresis hardware) DNA for experiments is linear and consists of a central “target” fragment, with labeled DNA fragments ligated at each end (see Fig. 2 for a sketch of the construct). The target fragment is typically on the order of a few kb as discussed earlier and is obtained from restriction-digested plasmid containing the sites and sequences of interest. The labeled DNA fragments used are
1 kb in length, incorporate either biotin or digoxigenin, and are obtained by PCR followed by restriction digestion and quantification. In the following we consider the use of XbaI and SacI sites. 7.1.1 Target DNA Plasmid containing the sequence of interest appropriately located between two restriction sites (e.g., XbaI and SacI) can readily be based on common high-copy-number plasmids such as pUC18:

Fig. 2 Sketch of DNA construct. A target DNA is digested with two restriction enzymes (RS1 and RS2) and ligated at one end to DNA bearing multiple biotin labels, and at the other end with DNA bearing multiple digoxigenin labels. The biotin-labeled DNA end can be anchored to a streptavidin-coated glass coverslip, and the digoxigenin-labeled end can be tethered to a magnetic bead coated with antidigoxigenin. Appropriate DNA sequence elements, such as a promoter to initiate transcription of DNA into RNA, can be positioned near the end of the DNA which is tethered to the glass surface so as to ensure the sequence is located within the TIR field. This enables, for instance, fluorescent imaging of labeled RNA polymerase which will bind to the promoter.

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1. Home-made competent DH5α bacteria are transformed with the plasmid using heat shock, and a single antibiotic-resistant colony is precultured in LB + ampicillin (typically, at 0.1 mg/mL final concentration) during the day, and used to inoculate an overnight culture of 250 mL of LB + ampicillin (0.1 mg/mL). 2. The next morning, cells are harvested and DNA extracted using alkaline lysis followed by gravity-flow ion-exchange chromatography (Macherey-Nagel NucleoBond Xtra Maxi Plus). 3. In the final steps of these protocols, DNA is precipitated with isopropanol, the precipitate is washed with 70% EtOH which is then evaporated off in a speedvac or at 37°C, and then resuspended, typically in 100–200 μL of Tris 10 mM pH 8. We note that centrifuging the resuspended DNA at 13,000  g for 10 min will pellet out any residual insoluble material, which may include carried-over resin or poorly resuspended DNA, for instance. The supernatant of this centrifugation step is thus typically devoid of any such potential aggregates. 4. DNA is quantified using a UV spectrophotometer (typically a few μg/μL are obtained). Its quality can be further ascertained by gel electrophoresis to determine the extent of nicked open, circular DNA in the preparation. In this procedure, a gel lacking any DNA stain (such as ethidium bromide) is used and the DNA revealed by staining only after the gel is run. One can thus quantify the fraction of plasmid which is supercoiled and hence unnicked. 5. 20 μg of plasmid DNA are then digested with the appropriate restriction enzymes; the target fragment of interest is obtained by purification on a 1% agarose gel and gel extraction on a spin column. DNA is resuspended in Tris 10 mM pH 8 and can in principle be stored for at least a year at 20°C without displaying a decline in the fraction of intact unnicked molecules as observed in the magnetic trap. 7.1.2 Labeled DNA 1. Using oligonucleotides Lambda-Forward (50 -gcg tat tag cga ccc atc gtc ttt ctg) and Lambda-Reverse (50 -gat gca cgc aat ggt gta gca ata att gc) as primers, and Lambda DNA as template, a PCR reaction incorporates dUTP-biotin or dUTP-digoxigenin into a
2.3-kbp PCR product.

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Lambda DNA (50 ng/mL)

1 μL

Lambda-Forward (10 μM)

7.5 μL

Lambda-Reverse (10 μM)

7.5 μL

dNTP mix (10 mM each)

25 μL

10 HiFidelity Buffer

25 μL

dUTP-biotin (1 mM) or dUTP-digoxigenin (1 mM)

5 μL

HiFi polymerase (3.5 U/μL)

5 μL

H2O

174 μL

A standard PCR program is used to amplify the DNA: 1. 95°C, 2 min. 2. 95°C, 15 s. 3. 57°C, 30 s. 4. 72°C, 1 min, 30 s. 5. Repeat steps 2–4, 25. 6. Hold at 16°C. 2. DNA is purified from the PCR reaction components using a standard glass membrane spin column (Macherey-Nagel NucleoSpin Gel & PCR Cleanup), taking care to remove carefully any residual ethanol from the column by pipetting, and/or gently drying off at 37°C following wash steps prior to DNA elution. 3. DNA is restriction digested with the appropriate enzyme (here XbaI and SacI, NEBiolabs ref. R0145 and R3156, respectively). The labeling scheme used here cleaves the DNA in the middle, and the efficiency of digestion can be determined using standard agarose gel electrophoresis to quantify the useful, cleaved fragment. The presence of biotin and dig moieties in the DNA can significantly reduce the efficiency of digestion, in a manner which depends on the extent to which thymines are present in the restriction site and the extent to which modified nucleotides are incorporated. 4. Target DNA and labeled DNA fragments are ligated for 3 h at room temperature:

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Target DNA (10 nM)

3 μL

Biotin-labeled DNA (100 nM)

1 μL

Digoxigenin-labeled DNA (100 nM)

1 μL

T4 DNA Ligase Buffer, 10

1 μL

H2O

3.5 μL

T4 DNA Ligase (NEBiolabs)

0.5 μL

5. The ligation reaction is terminated by heating to 65°C for 20 min. An aliquot of DNA from this reaction can be prepared for single-molecule experimentation by diluting the above reaction
60-fold, to
50 pM of target DNA.

8. PREPARATION OF ANTIDIGOXIGENINFUNCTIONALIZED MAGNETIC BEADS 8.1 Materials Dynabeads Tosylactivated MyOne magnetic beads (ThermoFisher Scientific, 65501) Sodium borate 0.1 M, pH 9.5 (SB buffer) Ammonium sulfate, 3 M, pH 9.5 Antidigoxigenin (Roche, 11 333 089 001) Rotating wheel maintained at 37°C BSA (Roche, 10 775 835 001) Tween-20 (Roche, 11 332 465 001) PBS, pH 7.4 1. 50 μL of Dynabeads Tosylactivated MyOne magnetic beads are washed with 100 μL of sodium borate buffer (SB) and resuspended in 10 μL of SB. 2. 200 μg of sheep polyclonal antidigoxigenin are then resuspended in 73.5 μL of SB and added to the 10 μL of washed and resuspended beads. We then add 41.5 μL of 3 M NH4SO4 (pH 9.5) to the bead/antibody mix and incubate for 24 h at 37°C on a rotating wheel to prevent sedimentation of the beads. 3. At the end of the incubation, the beads are pelleted and the supernatant discarded, and we add to the beads 125 μL of blocking buffer (1 PBS,

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pH 7.4, supplemented with 5 mg/mL BSA and 0.05% (w/v) Tween-20) (Roche). 4. The beads in blocking buffer are incubated overnight at 37°C on a rotating wheel, and the next day the beads are pelleted and the supernatant discarded. 5. The beads are then washed three times with 100 μL of washing/storage buffer (1 PBS, pH 7.4, supplemented with 1 mg/mL BSA and 0.05% (w/v) Tween-20) and resuspended in a final volume of 100 μL of washing/storage buffer. Beads are stored for about 1 month at 4°C.

9. ASSEMBLY OF BEAD-DNA SYSTEM AND LOADING OF REACTION CHAMBER 9.1 Materials Biotin- and dig-labeled DNA (see Labeled DNA from Section 5) Washing buffer (40 mM KHEPES, pH 8.0, 100 mM KCl, 8 mM MgCl2, 0.1 mg/mL BSA, 0.1% (w/v) Tween-20, 10 mM β-mercaptoethanol) Antidigoxigenin-derivatized polystyrene-coated glass surface (Section 6) And Dynabeads MyOne C1 streptavidin-coated magnetic beads (ThermoFisher, 65001) Or Streptavidin-derivatized PEGylated glass surface (Section 5) And Antidigoxigenin-derivatized magnetic beads (Section 8) 1. 5 μL of Dynabeads MyOne C1 streptavidin-coated magnetic beads are washed in 100 μL of washing buffer and resuspended in 5 μL of washing buffer.

9.1.1 Alternatively 1 μL of antidigoxigenin-functionalized magnetic beads are resuspended and washed in 10 μL of washing buffer, pelleted, and resuspended in 20 μL of washing buffer. 2. 1 μL of a 50 pM solution of DNA construct is placed at the bottom of a small microfuge tube.

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3. 5 μL of streptavidin-coated magnetic beads (alternatively, 2–3 μL of antidigoxigenin-coated beads) are mixed vigorously to the drop of DNA and allowed to incubate for 10–30 s. 4. 15 μL of washing buffer are added to the bead + DNA solution, preferably using a pipette tip which has been cut with a razor to provide a wider pipette bore and reduce shear forces on the bead + DNA system during pipetting. 5.
15 μL of the solution are injected into the flow cell by pipetting into one of the reservoirs glued at the end of the flow cell. Again, it is preferable to use a cut pipette tip for this. Beads can be observed by eye to enter the flow cell; it is recommended to allow beads to enter to only about 2/3 the length of the channel before allowing to rest
10–20 min (i.e., the time required for the beads to sediment to the surface and the DNA tether to the streptavidin). 6. Wash out unbound beads by flushing the flow cell with reaction buffer injected into the unused reservoir; typically
1 mL is used, in increments of
50 μL. It should be noted that excessively vigorous wash speeds can cause beads eventually to become stuck nonspecifically to the modified glass surface.

10. GENERAL CONSIDERATIONS FOR BUFFER PREPARATION Reusable filtration unit, polysulfone (Nalgene ThermoFisher, Waltham, MA, USA, DS0320-5045) Nylon membrane filters (Merck Millipore, GNWP04700) BSA (Roche, 10 775 835 001) Tween-20 (Roche, 11 332 465 001) Stock solutions are prepared from Milli-Q grade, UV-treated, 18.2 MΩ water, and reagents of at least molecular-biology grade purity. Stock solutions are filtered over nylon membrane filters using reusable polysulfonate filtration units. Enzymatic reactions contain ionic components as prescribed by biochemical assays and are nearly systematically supplemented with BSA (typically present at 0.1 mg/mL) and 0.1% (w/v) Tween-20 to minimize nonspecific interactions between biomolecules and surfaces. Buffers for single-molecule fluorescence should be verified to be fluorescence-free prior to use.

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11. CONCLUSIONS AND PERSPECTIVES Single-molecule experimentation continues to provide unique new tools for the study of biological and biophysical systems. Magnetic trapping provides a useful means for studying the mechanical properties of biomolecules, in particular the torsional and force-dependent properties of DNA, and also its interactions with proteins in real time. The development of robust correlative force spectroscopic and fluorescence methodologies will be invaluable for the study of complex multicomponent molecular systems. It will be exciting to watch future developments which can only include, one hopes, additional metrics such as electrical currents (Derrington et al., 2015).

ACKNOWLEDGMENTS This work was made possible by a EURYI grant, an EU 7th Framework Program grant HEALTH-F4-2008-223545, and a French ANR grant “RepOne,” in addition to core funding from the French National League Against Cancer, the CNRS, and the University of Paris Diderot. C.D. is supported by a PhD scholarship from the University of Paris Descartes and the Frontiers in Life Sciences-Bettencourt Doctoral Program, and J.F. is supported by a PhD scholarship from the China Scholarship Council.

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Gosse, C., & Croquette, V. (2002). Magnetic tweezers: Micromanipulation and force measurement at the molecular level. Biophysical Journal, 82, 3314–3329. Graves, E., Duboc, C., Fan, J., Stransky, F., Leroux-Coyau, M., & Strick, T. R. (2015). A dynamic DNA-repair complex observed by correlative single-molecule nanomanipulation and fluorescence. Nature Structural & Molecular Biology, 22, 452–457. Greenleaf, W. J., Frieda, K., Foster, D., Woodside, M. T., & Block, S. M. (2008). Direct observation of hierarchical folding in single riboswitch aptamers. Science, 319, 630–633. Hoskins, A., Friedman, L., Gallagher, S., Crawford, D., Anderson, E., Wombacher, R., et al. (2011). Ordered and dynamic assembly of single spliceosomes. Science, 331, 1289–1295. Howan, K., Smith, A. J., Westblade, L. F., Joly, N., Grange, W., Zorman, S., et al. (2012). Initiation of transcription-coupled repair characterized at single-molecule resolution. Nature, 490, 431–434. Huhle, A., Klaue, D., Brutzer, H., Daldrop, P., Joo, S., Otto, O., et al. (2015). Camera-based three-dimensional real-time particle tracking at kHz rates and a˚ngstr€ om accuracy. Nature Communications, 6, 5885. Ishijima, A., Kojima, H., Funatsu, T., Tokunaga, M., Higuchi, H., Tanaka, H., et al. (1998). Simultaneous observation of individual ATPase and mechanical events by a single myosin molecule during interaction with actin. Cell, 92, 161–171. Janovjak, H., Kessler, M., Oesterhelt, D., Gaub, H. E., & M€ uller, D. J. (2003). Unfolding pathways of native bacteriorhodopsin depend on temperature. The EMBO Journal, 22, 5220–5229. Kapanidis, A. N., Margeat, E., Ho, S., Kortkhonjia, E., Weiss, S., & Ebright, R. H. (2006). Initial transcription by RNA polymerase proceeds through a DNA-scrunching mechanism. Science, 314, 1144–1147. Kapanidis, A. N., & Strick, T. R. (2009). Biology, one molecule at a time. Trends in Biochemical Sciences, 34, 234–243. Lang, M. J., Fordyce, P. M., Engh, A. M., Neuman, K. C., & Block, S. M. (2004). Simultaneous, coincident optical trap and single-molecule fluorescence. Nature Methods, 1, 133–139. Lionnet, T., Allemand, J.-F., Revyakin, A., Strick, T. R., Saleh, O. A., Bensimon, D., et al. (2012a). Magnetic trap construction. Cold Spring Harbor Protocols, 2012, 133–138. Lionnet, T., Allemand, J.-F., Revyakin, A., Strick, T. R., Saleh, O. A., Bensimon, D., et al. (2012b). Single-molecule studies using magnetic traps. Cold Spring Harbor Protocols, 2012, 34–49. Lipfert, J., Wiggin, M., Kerssemakers, J. W., Pedaci, F., & Dekker, N. H. (2011). Freely orbiting magnetic tweezers to directly monitor changes in the twist of nucleic acids. Nature Communications, 2, 439. Myong, S., Bruno, M., Pyle, A. M., & Ha, T. (2007). Spring-loaded mechanism of DNA unwinding by hepatitis C virus NS3 helicase. Science, 317, 513–516. Rief, M., Gautel, M., Schemmel, A., & Gaub, H. E. (1998). The mechanical stability of immunoglobulin and fibronectin III domains in the muscle protein titin measured by atomic force microscopy. Biophysical Journal, 75, 3008–3014. Robb, N. C., Cordes, T., Hwang, L. C., Gryte, K., Duchi, D., Craggs, T. D., et al. (2013). The transcription bubble of the RNA polymerase-promoter open complex exhibits conformational heterogeneity and millisecond-scale dynamics: Implications for transcription start-site selection. Journal of Molecular Biology, 425, 875–885. Sellers, J., & Veigel, C. (2010). Direct observation of the myosin-Va power stroke and its reversal. Nature Structural and Molecular Biology, 17, 590–595. Smith, S. B., Finzi, L., & Bustamante, C. (1992). Direct mechanical measurements of the elasticity of single DNA molecules by using magnetic beads. Science, 258, 1122–1126. Strick, T. R., Allemand, J.-F., Bensimon, D., Bensimon, A., & Croquette, V. (1996). The elasticity of a single supercoiled DNA molecule. Science, 271, 1835–1837.

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CHAPTER TWELVE

Measuring Force-Induced Dissociation Kinetics of Protein Complexes Using Single-Molecule Atomic Force Microscopy K. Manibog*,†,1, C.F. Yen*,†,1, S. Sivasankar*,†,2 *Iowa State University, Ames, IA, United States † Ames Laboratory, U.S. Department of Energy, Ames, IA, United States 2 Corresponding author: e-mail address: [email protected]

Contents 1. Introduction 2. Models for the Mechanical Response of Receptor–Ligand Bonds 2.1 Slip Bonds 2.2 Catch Bonds 2.3 Ideal Bonds 3. Measuring in vitro Force-Dependent Kinetics With an AFM 3.1 Principles of AFM Operation 3.2 Engineering Proteins for AFM Force Measurements 3.3 Surface Functionalization 3.4 Force Clamp Measurement 3.5 Data Analysis 4. Using AFM Force Measurements to Characterize in vivo Unbinding Kinetics 4.1 Possible Experimental Configurations 4.2 Sample Preparation 4.3 Force Measurement 4.4 Analyzing the Force vs Distance Traces 5. Limitations of Current Technologies and Future Directions Acknowledgments References

298 299 299 300 301 301 301 304 304 305 307 310 310 312 312 313 314 316 316

Abstract Proteins respond to mechanical force by undergoing conformational changes and altering the kinetics of their interactions. However, the biophysical relationship between mechanical force and the lifetime of protein complexes is not completely understood. In this chapter, we provide a step-by-step tutorial on characterizing the force1

Contributed equally to this work.

Methods in Enzymology, Volume 582 ISSN 0076-6879 http://dx.doi.org/10.1016/bs.mie.2016.08.009

#

2017 Elsevier Inc. All rights reserved.

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dependent regulation of protein interactions using in vitro and in vivo single-molecule force clamp measurements with an atomic force microscope (AFM). While we focus on the force-induced dissociation of E-cadherins, a critical cell–cell adhesion protein, the approaches described here can be readily adapted to study other protein complexes. We begin this chapter by providing a brief overview of theoretical models that describe force-dependent kinetics of biomolecular interactions. Next, we present step-by-step methods for measuring the response of single receptor–ligand bonds to tensile force in vitro. Finally, we describe methods for quantifying the mechanical response of single protein complexes on the surface of living cells. We describe general protocols for conducting such measurements, including sample preparation, AFM force clamp measurements, and data analysis. We also highlight critical limitations in current technologies and discuss solutions to these challenges.

1. INTRODUCTION Mechanical signals play essential roles in diverse physiological and pathological processes such as promoting cancer metastasis (Wirtz, Konstantopoulos, & Searson, 2011) and stem cell differentiation (Engler, Sen, Sweeney, & Discher, 2006). In general, mechanical forces trigger protein conformational changes which in turn lead to the exposure of cryptic ligandbinding sites (del Rio et al., 2009), enzymatic cleavage sites (Gumpp et al., 2009), or to strengthened receptor–ligand interactions (Hertig & Vogel, 2012; Rakshit & Sivasankar, 2014; Zhu, 2014). However, the relationship between force and receptor–ligand dissociation is not completely understood. In this chapter, we provide a step-by-step tutorial on characterizing the forcedependent dissociation of protein complexes using in vitro and in vivo singlemolecule force measurements with an atomic force microscope (AFM). While we focus on the mechanical response of E-cadherin (Ecad), an essential homotypic cell–cell adhesion protein, the approaches described here can be readily adapted to study the other receptor–ligand bonds in vitro and in vivo. Ecads are transmembrane proteins; while their extracellular region interacts with Ecads on opposing cells to mediate cell–cell adhesion, their cytoplasmic region connects to the cytoskeleton via a series of effector proteins that assist in signal transduction. Ecads enable multicellular assemblies to withstand mechanical force and are essential in tissue formation and in maintaining tissue integrity (Baker et al., 2015; Hahn & Schwartz, 2009; Leckband & de Rooij, 2014). The kinetics of Ecad interactions are tuned via both physical factors like mechanical force and cellular factors such as the formation of cell-surface clusters and interaction with cytoplasmic regulatory proteins. In this chapter, we present how AFM-based methods can be used to characterize the interplay of force and cellular factors in regulating the kinetics of Ecad binding.

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2. MODELS FOR THE MECHANICAL RESPONSE OF RECEPTOR–LIGAND BONDS When receptors and ligands interact, they assume their most energetically favorable conformation. Receptor–ligand complexes dissociate by overcoming a confining energy of barrier of height, △E; under equilibrium conditions, the dissociation rate, k0off is exponentially related to ΔE and is proportional to the frequency of attempts, A to cross this barrier:
 ΔE 0 koff ¼ A exp (1) kB T where kB is the Boltzmann constant and T is the temperature. Tensile stress alters the height of the energy barrier thereby changing the rate of receptor– ligand dissociation. Based on their response to tensile force, receptor–ligand interactions can be classified into one of three types of bonds: slip bonds, catch bonds, and ideal bonds (Fig. 1) (Rakshit & Sivasankar, 2014).

2.1 Slip Bonds Slip bonds are conventional interactions where pulling force accelerates the rupture of the receptor–ligand complex (Fig. 1A). As first proposed

Fig. 1 Slip, catch-slip, and ideal bonds. (A) A slip bond weakens in the presence of force while (B) a catch-slip bond initially strengthens and subsequently weakens when pulled. (C) An ideal bond is insensitive to tensile stress. All panels in this figure were measured using AFM force clamp spectroscopy on recombinant Ecad mutants trapped in different conformations. The data in panels (A) and (C) are from Manibog, K., Sankar, K., Kim, S., Zhang, Y., Jernigan, R. L., & Sivasankar, S. (2016). Molecular determinants of cadherin ideal bond formation: Conformation dependent unbinding on a multidimensional landscape. Proceedings of the National Academy of Sciences of the United States of America, 113(39), E5711–E5720. http://dx.doi.org/10.1073/pnas.1604012113. Panel (B) is adapted from Manibog, K., Li, H., Rakshit, S., & Sivasankar, S. (2014). Resolving the molecular mechanism of cadherin catch bond formation. Nature Communications, 5, 3941. http://dx.doi. org/10.1038/ncomms4941.

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by Bell (1978), the lifetime of the bond, τ(F) decreases exponentially with force, F as:
 FΔx τðF Þ ¼ τ0 exp  (2) kB T where τ0 is the intrinsic bond lifetime and Δx is the distance between the bound and transition states along the pulling coordinate. This theory assumes that Δx is positive and that ΔE is high enough that force does not alter the position of the transition state. As a consequence, increasing the force lowers the height of the energy barrier by ΔE  FΔx (cf. Eq. 1) resulting in an exponential decrease of bond lifetimes. This simple theory has been expanded to include the force-free activation energy and the shape of the free-energy profile (Dudko, Hummer, & Szabo, 2006).

2.2 Catch Bonds Catch bonds resist dissociation when subjected to a mechanical force; their lifetimes increase with increasing force application (Fig. 1B). Although catch bonds were first directly measured relatively recently for the dissociation of the adhesion molecule P-selectin and its ligand (Marshall et al., 2003), numerous proteins such as integrins (Kong, Garcı´a, Mould, Humphries, & Zhu, 2009), FimH (Thomas, Trintchina, Forero, Vogel, & Sokurenko, 2002), vWF (Yago et al., 2008), and Ecads (Manibog, Li, Rakshit, & Sivasankar, 2014; Rakshit, Zhang, Manibog, Shafraz, & Sivasankar, 2012) have now been reported to exhibit catch bond behavior. Currently, there is no unified model that can describe catch bonds formed by different receptor–ligand complexes. Therefore, several theoretical models have been proposed to describe the process by which receptor–ligand complexes form catch bonds. Most of these models including the one-bound state two pathway model (Pereverzev, Prezhdo, Forero, Sokurenko, & Thomas, 2005), two-bound state two pathway model (Barsegov & Thirumalai, 2005), single dissociation pathway in a multidimensional landscape model (Suzuki & Dudko, 2010), and fluctuating energy landscape model (Liu & Ou-Yang, 2006) are energy landscape based. There are also a few structural models such as the direct deformation model (Pereverzev & Prezhdo, 2006), allosteric deformation models (Pereverzev, Prezhdo, & Sokurenko, 2009), and sliding–rebinding model (Lou & Zhu, 2007), which propose structural deformations that result in catch bond formation. The biphasic catch-slip bond exhibited by Ecad is best described by the sliding–rebinding model (Lou & Zhu, 2007) where external force induces opposing protomers to slide past

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each other and rebind in a new conformation with additional force-induced interactions that increase bond lifetimes (Fig. 1B).

2.3 Ideal Bonds Ideal bonds are force–insensitive interactions with lifetimes that are independent of applied force (Fig. 1C). Ideal bonds are a special case of a slip bond, when the interaction energies of the receptor ligand complex are harmonic in nature, exhibiting equal stiffness, and resting lengths in the bound state and transition state (Dembo, Torney, Saxman, & Hammer, 1988). Ideal bonds are also possible when biomolecules unbind in a multidimensional landscape where the extensions of the bound and the transition states are identical along the pulling direction (Manibog et al., 2016; Suzuki & Dudko, 2010). The first experimental observation of ideal bonds was described in our research with Ecads (Rakshit et al., 2012).

3. MEASURING IN VITRO FORCE-DEPENDENT KINETICS WITH AN AFM AFM is a scanning probe microscope technique that was initially developed to image and characterize the properties of surfaces with subnanometer resolution (Binnig, Quate, & Gerber, 1986). However, the development of techniques for functionalizing and decorating probes and substrates with biomolecules have made AFM an ideal tool for biological investigations, in particular for studying the mechanical properties of biomolecular complexes in the presence of external applied force (Neuman & Nagy, 2008). An advantage of AFM is its high spatial and temporal resolution and its ability to interrogate single molecules under near physiological conditions. In addition, the results of AFM force measurements can be compared with protein interaction studies using computational methods (Lou & Zhu, 2007; Manibog et al., 2014). In this section, we will provide a brief introduction to AFM-based experiments to characterize the lifetimes of receptor–ligand bonds at the single-molecule level.

3.1 Principles of AFM Operation AFM can be used to monitor the unbinding or unfolding of single molecules in the presence of either a constant rate of force application (force ramp) or a constant force (force clamp). A force ramp measurement is implemented when one wishes to determine the effect of loading rates on the kinetics of protein unbinding, while AFM force clamp is employed to directly measure the rates of unbinding or unfolding under a constant tensile force

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(Evans & Calderwood, 2007). The in vitro measurement of force-dependent kinetics of receptor–ligand interactions is typically performed using AFM force clamp experiments. In order to perform an AFM force clamp measurement, the AFM force probe and substrate are first functionalized with biomolecules of interest. The force probe is typically a thin silicon or silicon nitride cantilever that acts as a simple harmonic spring. Either the functionalized AFM cantilever or the sample is translated using a piezoelectric actuator (PZT) that can operate in either open-loop or closed-loop modes (Fig. 2A). The functionalized AFM cantilever and substrate are first brought into contact to enable receptor–ligand interaction. The cantilever is then separated from the substrate to load the receptor–ligand complex. Changes in force due to receptor–ligand loading result in a deflection of the cantilever which is measured by reflecting a focused laser beam off the backside of the cantilever onto a position-sensitive quadrant photodetector (QPD). The spring constant, k of the cantilever is determined by monitoring its thermal fluctuations when the cantilever is far away from the surface such that it vibrates with amplitude, z0 around its equilibrium position at temperature, T (Hutter & Bechhoefer, 1993). From the equipartition theorem, the

Fig. 2 Schematic of AFM force measurement and sensitivity calibration. (A) Schematic of setup used in single-molecule AFM force clamp measurement. The AFM cantilever is mounted on a PZT scanner, which controls cantilever movement. The tip and substrate are functionalized with PEG linkers, some of which are decorated with streptavidins. Biotinylated Ecads are attached to the streptavidins. (B) Representative QPD detector voltage (V) vs PZT movement during sensitivity calibration. Calibration is performed by pressing the AFM cantilever on a hard substrate, which results in cantilever deflection (increase in V). The measured sensitivity is calculated from the slope.

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thermal energy is approximately equal to the average vibrational energy due to cantilever oscillations:   1 2 2 1 (3) mω z0 ¼ kB T 2 2 where m is the effective mass of the cantilever, ω is the resonant frequency, and kB is the Boltzmann constant. Since the resonant frequency is related to pffiffiffiffiffiffiffiffi the spring constant by ω ¼ k=m, the spring constant can be obtained as   k ¼ kB T = z0 2 . From the computed spring constant of the cantilever, the cantilever deflection is converted to force using Hooke’s Law: F ¼ kΔx

(4)

where the cantilever deflection Δx ¼ ΔV =S where ΔV is the voltage difference on the QPD due to cantilever deflection and S is the optical lever sensitivity (voltage change per unit distance (V/nm)). The displacement between the AFM tip and the underlying substrate, Δz, is measured as Δz ¼ ΔD  Δx where ΔD is the PZT displacement. S is typically measured by using the PZT to press the cantilever on a hard, nondeformable surface and then moving it by a known displacement, ΔD1, while simultaneously monitoring the voltage change (ΔV1) on the QPD (Fig. 2B). The sensitivity is calculated as: S ¼ ΔV1 =ΔD1 . Since position sensors in a closed-loop PZT are used to ensure accurate displacement when the actuator is driven by an input voltage, the motion of a closed-loop PZT is linear and S is measured accurately. In contrast, hysteresis and creep behaviors result in nonlinear displacement when a PZT is operated in open-loop mode (Hall, 2001; Lapshin, 1995). These nonlinearities can contribute to substantial errors in piezo movement, increasing errors in measured sensitivity. Accurate values of measured sensitivity are critical for determining interaction forces, calculating cantilever spring constants, and calculating the tip–substrate distance (Butt, Cappella, & Kappl, 2005). An important factor to consider in AFM experiments is that small cantilever oscillations caused by mechanical vibrations, acoustic noise, air turbulence, and thermal drift decrease the signal-to-noise ratio. Low frequency mechanical vibrations can be reduced by placing the AFM system on active or passive isolation supports such as optical table or bungee cord isolators. Enclosing the system inside an isolation box with layers of acoustic-damping materials decreases the effect of acoustic noise or air turbulence. The effect of thermal drift can be minimized by stripping the reflective metal coating off

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the backside of the cantilever (Churnside et al., 2012) or by allowing the AFM to “settle” after loading an AFM cantilever.

3.2 Engineering Proteins for AFM Force Measurements Recombinant proteins can be generated for AFM force measurements using standard molecular biology techniques. The only requirement is that the proteins should be engineered with a molecular handle that can be used to immobilize the protein on a surface. While a wide variety of immobilization chemistries such as immobilization via Cys residues (Dietz et al., 2006) have been described in the literature, we use streptavidin–biotin chemistry to immobilize Ecad in our experiments. Briefly, the full-length extracellular domains of Ecad with a C-terminal Avi-tag (for biotinylation), a Tev sequence (for proteolytic cleavage), and a His-tag (for protein purification) is expressed in HEK293T cells, and purified from the conditioned media using a Nickel NTA resin. After protein purification, the Avi-tag sequence is biotinylated using BirA enzyme (BirA500 kit; Avidity). Since these protocols have been extensively described in our previous works (Manibog et al., 2014; Rakshit et al., 2012; Zhang, Sivasankar, Nelson, & Chu, 2009) and are not the main focus of this review, we will not discuss these methods in greater detail.

3.3 Surface Functionalization While the purified recombinant protein can be immobilized on AFM cantilevers and glass coverslips using a range of chemistries (Dietz et al., 2006; Popa, Kosuri, Alegre-Cebollada, Garcia-Manyes, & Fernandez, 2013; Yu, Malkova, & Lyubchenko, 2008), it is important that the proteins are attached to polyethylene glycol (PEG) tethers organized as dense monolayers on the cantilever and coverslip. Specifically immobilizing the protein on flexible PEG linkers serves three purposes (i) functionalizing cantilevers and coverslips with a dense PEG cushion reduces the nonspecific adsorption of protein; (ii) flexible PEG tethers enable unhindered interactions between specifically bound proteins during tip–substrate encounters; and (iii) the stretching of PEG serves as a molecular fingerprint for single-molecule unbinding since its extension under load has been extensively characterized (Friedsam, Wehle, Kuhner, & Gaub, 2003; Oesterhelt, Rief, & Gaub, 1999). The first step in functionalizing the AFM cantilevers and coverslips is to clean them by soaking them overnight in a 25% H2O2:75% H2SO4 solution at room temperature and then washing them with deionized (DI) water.

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The coverslips are cleaned further by sequential sonication in 1 M KOH and DI water for 10 min. Finally, the coverslips and cantilevers are sonicated three times in 99.5% grade acetone; each sonication step lasts for 5 min. The cleaned cover slips and cantilevers are functionalized with amine groups by incubating them for 30 min in 2% (v/v) solution of 3-aminopropyltriethoxysilane (Sigma) dissolved in acetone. The free amine groups on the silanized cantilevers and coverslips are decorated with PEG tethers (Laysan Bio) that contains an amine-reactive N-hydroxysuccinimide ester at one end. Depending on the immobilization chemistry that is employed, the functional group on the free-end of the immobilized PEG tether can be varied. For instance, we recently covalently immobilized recombinant prion proteins with Cys residues on glass substrates and AFM cantilevers decorated with maleimide functionalized PEG tethers (Yen, Harischandra, Kanthasamy, & Sivasankar, 2016). We also routinely immobilize biotinylated Ecads on coverslips and cantilevers that are functionalized with biotinylated PEG tethers decorated with streptavidins. In both these cases, anywhere from 5% to 10% of the PEG spacers contain biotin/maleimide groups at their free-end, while the remaining PEGs are inert. The PEG-functionalized cantilevers can be stored in a vacuum desiccator and used for force measurement within 2 weeks. Prior to beginning an Ecad experiment, a PEG-functionalized coverslip and AFM cantilever are incubated in 0.1 mg/mL BSA for approximately 12 h to minimize nonspecific protein binding. The substrate and cantilever are then incubated with 0.1 mg/mL streptavidin (Sigma) for 30 min. Finally, the coverslips and cantilevers are decorated with Ecads by incubating them for 45 min with 100–200 nM biotinylated Ecads (Fig. 2A). Following Ecad immobilization, free biotin-binding sites on the streptavidins are blocked using 10-min incubation with 2 μM biotin; the free biotins are flushed away after blocking. A pH 7.5 buffer solution (10 mM Tris, 100 mM NaCl, 10 mM KCl, and 2.5 mM CaCl2) is used in all incubation and subsequent measurement steps.

3.4 Force Clamp Measurement A typical AFM force clamp experiment begins with the functionalized cantilever in an equilibrium position far from the functionalized substrate (Fig. 3). The cantilever is then brought into contact with the substrate and gently pressed allowing the proteins to interact and form bonds. It is

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Fig. 3 Typical AFM force clamp measurement of an Ecad–Ecad complex. The Ecadfunctionalized substrate and AFM tip are first brought into contact. The tip is pressed against the surface at a force of 80 pN and held for 3 s. The AFM tip is then withdrawn from the surface and “clamped” at a predetermined force so that a constant pulling force is applied to the Ecad–Ecad bond. Once the bond breaks, the cantilever recoils to its equilibrium position. Survival time at the clamping force is determined from the period of time that the Ecad-dimer persists. This figure is adapted from Rakshit, S., Zhang, Y., Manibog, K., Shafraz, O., & Sivasankar, S. (2012). Ideal, catch, and slip bonds in cadherin adhesion. Proceedings of the National Academy of Sciences of the United States of America, 109(46), 18815–18820. http://dx.doi.org/10.1073/pnas.1208349109.

important to note that the pressing force should be minimized (90% of the measurements corresponds to single-molecule events. Upon retracting the AFM cantilever from the substrate, a force is applied to the bonds formed between Ecads on the cantilever and the cell. Similar to in vitro measurements, the applied force can be maintained by keeping the QPD deflection signal constant using a closed-loop feedback. Alternatively, and more simply, a membrane tether (formed by pulling the membrane bound Ecad) can be used to exert a constant force on the bond (Krieg, Helenius, Heisenberg, & Muller, 2008). The physical model of membrane flow predicts that the force required to extract a membrane tether is positively correlated to the pulling velocity (Hochmuth, Wiles, Evans, & McCown, 1982; Krieg et al., 2008). Assuming that the composition and physical properties of the membrane remain unchanged, the Ecad-dimer can be clamped at a constant force by simply retracting the cantilever at a constant speed. Increasing the pulling velocity leads to a stronger clamp force (Fig. 5B). Taking advantage of the membrane tethering effect, the in vivo force clamp experiment can be simplified into the following three steps: (i) move the AFM cantilever toward the substrate until the QPD deflection signal increases by 0.1 to 0.5 V (depends on the desired contact force); (ii) Hold the cantilever on the cell surface for a predefined contact time (typically in the range of 0.1–10 s); (iii) Withdraw the cantilever from the substrate at a constant speed (typically in the range of 0.5–20 μm/s). To make the results statistically reliable, the measurements should be repeated at different positions over several cells and the substrate should be replaced every 4–6 h, before the cells start detaching.

4.4 Analyzing the Force vs Distance Traces A force–distance curve obtained using an Ecad-functionalized cantilever and cell-coated substrate with a contact force of 250 pN is shown in Fig. 5C. The QPD deflection signal is converted into force using the same protocol described in the in vitro data analysis (Section 3.1). Only data recorded during the cantilever retraction are analyzed. Due to interactions between the Ecads on the AFM stylus and the cell surface, membrane tethers are formed; rupture of these interactions results in a “step-like” decrease in the force–distance curve (Fig. 5C). To correct for force changes caused by hydrodynamic drag and thermal/mechanical drift, data after the final force drop (labeled “drop” in Fig. 5C) are fitted to a straight line and used for baseline correction (red line in Fig. 5C).

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Ideally, most force–distance traces show a single membrane tether stretching event (Fig. 5C), indicating that the measured data originate from the rupture of a single molecular bond. The likelihood of obtaining single membrane tether stretching data can be enhanced by decreasing the contact force and interaction time. However, since the contact force cannot be accurately controlled (due to the compliance of the cell surface), the force traces often show several membrane tether stretching events. When multiple membrane tethers are pulled in parallel (inset, Fig. 5C), the force distribution across the multiple bonds is uncertain (Friedrichs et al., 2013). Therefore, only the final membrane tether stretching should be used in subsequent data analysis. Furthermore, given that both specific unbinding events (force drops due to the rupture of Ecad–Ecad interaction) and nonspecific unbinding events can be measured, it is important to quantify the fraction of nonspecific adhesion events by using cantilevers without the receptor and also by performing experiments in the presence of function blocking antibodies/ligands. Since the probe is withdrawn at a constant speed, bond lifetime can be simply calculated by dividing the rupture distance (“d1” in Fig. 5C) by the retraction speed. The height of force step can be determined from the difference of average forces before and after the force drop. More sophisticated algorithms based on Chi-square statistics or step-finding algorithms can also be applied for automated step detection (Kerssemakers et al., 2006; Sariisik et al., 2013). Identical to the data analysis for in vitro force clamp experiments, lifetimes for different clamping forces are grouped and converted into survival probability plots (Fig. 5D). The mean bond lifetimes at different clamping forces can be obtained by fitting the corresponding survival probability distributions to a single exponential decay. Our preliminary results show that Ecad bond lifetimes measured in vivo are approximately 50–100 times higher than in vitro lifetimes. Possible reasons for the enhanced in vivo Ecad bond lifetimes are (i) formation of Ecads clusters on the cell surface (Wu et al., 2015) that interact as a single adhesive unit with the Ecads on the AFM tip and (ii) inside-out signaling by cytoplasmic effectors bound to the Ecad intracellular region.

5. LIMITATIONS OF CURRENT TECHNOLOGIES AND FUTURE DIRECTIONS Single-molecule AFM force clamp spectroscopy is a highly versatile technique that has been used to characterize a wide range of receptor–ligand complexes. However, several technical drawbacks limit the widespread adoption of this technique. Slow data acquisition rates of commercial

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AFM systems limit the temporal resolution of bond lifetime measurements to several hundreds of μs. However, recent advances in the development of high-speed AFMs (Ando, Uchihashi, & Kodera, 2013) and in cantilever design (Edwards et al., 2015) are likely to significantly enhance the temporal resolution of AFM force clamp spectroscopy measurements. A second limitation of single-molecule force clamp measurements is the ambiguity involved in discriminating specific single-molecule unbinding events and nonspecific interactions. While the data analysis described in Section 3.5 is useful in identifying specific unbinding events, this approach is still lacking. Similarly, while the unfolding of polyprotein constructs are commonly used as molecular fingerprints in AFM force clamp measurements of protein unfolding (Liang & Fernandez, 2009), they cannot be used in receptor–ligand unbinding experiments due to the high forces required to unfold a protein domain. Consequently, new chemistries that minimize nonspecific protein adsorption, enhance specific binding, and serve as low-force molecular fingerprints are essential. Although force clamp spectroscopy is powerful in resolving singlemolecule interactions on living cells, it is limited by time consuming data collection and sample preparation methods. The requirement of home-built programs for instrument control and data analysis has also slowed the popularization of this technique. Fortunately, AFM modules that have been specifically designed for the investigation of cell adhesion are now commercially available. A main challenge in live cell force measurements lies in data interpretation since every measurement is the product of several cellular behaviors, including specific protein interactions, compositional change of membrane lipids, cellular elasticity, cytoskeleton dynamics, and nonspecific adhesions. Correlating the force trace features to specific molecular interactions is a major thrust in the field. Finally, while single-molecule AFM force spectroscopy measurements have proven very powerful in determining the kinetics of receptor–ligand interactions, they do not always report on mechanically induced changes in structure. These conformational changes can be monitored using complementary fluorescence-based techniques such as fluorescence resonance energy transfer (FRET). While other force measurement techniques like optical tweezers have been successfully integrated with single-molecule fluorescence and FRET by several groups (Comstock, Ha, & Chemla, 2011; Hohng et al., 2007; Lang, Fordyce, Engh, Neuman, & Block, 2004; Tarsa et al., 2007), recent attempts at combining AFM and FRET (He, Lu, Cao, & Lu, 2012) have not been widely adopted by the single-molecule

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biophysics community due to the low throughput of the instrument. Despite this limitation, researchers have used combined single-molecule AFM and fluorescence-based approaches to track force-induced protein unfolding (Sarkar, Robertson, & Fernandez, 2004), to monitor force-induced enhancement of enzymatic catalysis (Gumpp et al., 2009), to monitor force-induced conformational changes in a single protein (He et al., 2012), and to “cut and paste” single fluorescent DNA and protein molecules for the bottom-up assembly of nanoscale structures (Kufer, Puchner, Gumpp, Liedl, & Gaub, 2008; Puchner, Kufer, Strackharn, Stahl, & Gaub, 2008; Strackharn, Pippig, Meyer, Stahl, & Gaub, 2012). Building an instrument capable of high throughput, automated, simultaneous single-molecule force, and fluorescence measurements is currently an active area of research.

ACKNOWLEDGMENTS This research was supported in part by grants from the American Heart Association (Scientist Development Grant 12SDG9320022), from the American Cancer Society (Research Scholar Grant 124986-RSG-13-185-01-CSM), and from the National Science Foundation (PHY-1607550).

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CHAPTER THIRTEEN

Improved Force Spectroscopy Using Focused-Ion-BeamModified Cantilevers J.K. Faulk*,1, D.T. Edwards*,1, M.S. Bull*, T.T. Perkins*,†,2 *JILA, National Institute of Standards and Technology and University of Colorado, Boulder, CO, United States † University of Colorado, Boulder, CO, United States 2 Corresponding author: e-mail address: [email protected]

Contents 1. Introduction 2. Overview of Modification Process 3. Methods and Protocols 3.1 Equipment and Materials 3.2 Mounting and Loading AFM Cantilevers 3.3 Imaging with the SEM and Preparing the FIB 3.4 Modifying BioLever Minis with the FIB 3.5 Depositing TEOS on BioLever Minis 3.6 Etching of BioLever Minis 3.7 Modifying BioLever Fasts with the FIB 3.8 Characterizing Cantilever Performance 4. Improved Performance of FIB-Modified Cantilevers 5. Conclusions Acknowledgments References

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Abstract Atomic force microscopy (AFM) is widely used in biophysics, including forcespectroscopy studies of protein folding and protein–ligand interactions. The precision of such studies increases with improvements in the underlying quality of the data. Currently, data quality is limited by the mechanical properties of the cantilever when using a modern commercial AFM. The key tradeoff is force stability vs short-term force precision and temporal resolution. Here, we present a method that avoids this compromise: efficient focused-ion-beam (FIB) modification of commercially available cantilevers. Force precision is improved by reducing the cantilever’s hydrodynamic drag, and force stability is improved by reducing the cantilever stiffness and by retaining a cantilever’s 1

These authors contributed equally to this manuscript.

Methods in Enzymology, Volume 582 ISSN 0076-6879 http://dx.doi.org/10.1016/bs.mie.2016.08.007

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gold coating only at its free end. When applied to a commonly used short cantilever (L ¼ 40 μm), we achieved sub-pN force precision over 5 decades of bandwidth (0.01–1000 Hz) without significantly sacrificing temporal resolution (75 μs). Extending FIB modification to an ultrashort cantilever (L ¼ 9 μm) also improved force precision and stability, while maintaining 1-μs-scale temporal resolution. Moreover, modifying ultrashort cantilevers also eliminated their inherent underdamped high-frequency motion and thereby avoided applying a rapidly oscillating force across the stretched molecule. Importantly, fabrication of FIB-modified cantilevers is accessible after an initial investment in training. Indeed, undergraduate researchers routinely modify 2–4 cantilevers per hour with the protocol detailed here. Furthermore, this protocol offers the individual user the ability to optimize a cantilever for a particular application. Hence, we expect FIBmodified cantilevers to improve AFM-based studies over broad areas of biophysical research.

ABBREVIATIONS AFM atomic force microscopy FIB focused-ion-beam PSD power spectral density SEM scanning electron microscopy SMFS single-molecule force spectroscopy TEOS tetraethyl orthosilicate RSA reduced-scan area HFW horizontal field width QPD quadrant photodiode

1. INTRODUCTION Atomic force microscopy (AFM) is a powerful technique used in diverse scientific fields, ranging from nanoscience to biophysics, due to its ability to image with sub-nm lateral resolution (Muller & Dufrene, 2008). A complementary application of AFM is single-molecule force spectroscopy (SMFS) (Greenleaf, Woodside, & Block, 2007; Neuman & Nagy, 2008), such as the mechanical unfolding of individual proteins (Rief, Gautel, Oesterhelt, Fernandez, & Gaub, 1997) and disruption of single protein– ligand bonds (Florin, Moy, & Gaub, 1994; Lee, Kidwell, & Colton, 1994). The precision with which AFM can be applied to any of these applications is enhanced by improving the quality of the underlying data. Currently, choosing a cantilever for a measurement necessarily requires sacrificing in at least one of three key metrics: force precision, force stability, or temporal resolution. However, by modifying the nanomechanical

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Fig. 1 Improving force spectroscopy with focused-ion-beam (FIB)-modified cantilevers. (A) An FIB-modified BioLever Mini, and (B) an FIB-modified BioLever Fast. At the top left of each column, scanning electron microscopy (SEM) images of the cantilever and its stiffness are shown. At the top right of each column, the response time of each cantilever to an abrupt change in F is shown. Time constants were determined by an exponential fit (dashed line) to each record. For the FIB-modified BioLever Mini, the traces shown are the averaged step-response function during protein unfolding recorded at 50 kHz. For the FIB-modified BioLever Fast, the record shown is the response to detachment of the polyprotein from the tip recorded at 5 MHz. SEM image in panel (A) is reprinted with permission from Bull M. S., Sullan, R. M., Li, H., & Perkins, T. T. (2014). Improved single molecule force spectroscopy using micromachined cantilevers. ACS Nano, 8, 4984–4995. SEM image in panel (B) is reprinted with permission from Edwards D. T., Faulk, J. K., Sanders, A. W., Bull, M. S., Walder, R., LeBlanc, M. A., et al. (2015). Optimizing 1-μsresolution single-molecule force spectroscopy on a commercial atomic force microscope. Nano Letters, 15, 7091–7098.

properties of the cantilever, we obtain cantilevers that offer a significantly better combination of these performance metrics over commercially available cantilevers (Fig. 1) (Bull, Sullan, Li, & Perkins, 2014; Edwards et al., 2015; Edwards & Perkins, 2016). This chapter details how to use focusedion-beam (FIB) milling to reduce the stiffness and the hydrodynamic drag of two commonly used cantilevers in conjunction with removing the cantilever’s gold coating from the majority of its surface area to achieve state-ofthe-art performance for AFM-based SMFS. Historically, AFM was the instrument of choice when mechanically unfolding single proteins (Carrion-Vazquez et al., 1999; Rief et al., 1997). More recently, optical-trapping-based assays have been used to probe the equilibrium folding and unfolding of proteins under force due to the better force stability and short-term force precision of optical traps relative to

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AFM (Cecconi, Shank, Bustamante, & Marqusee, 2005; Stigler, Ziegler, Gieseke, Gebhardt, & Rief, 2011; Yu et al., 2012). Force stability is important in SMFS as unfolding rates are exponentially sensitive to changes in F. Improved force precision enables detecting subtle or rapid changes in molecular structure. Despite AFM’s historical weaknesses in force stability and force precision relative to optical traps, AFM remains a popular technique as stable instruments are commercially available. Such AFMs provide dramatically enhanced ease of use relative to custom-built optical traps, broadening the range of potential users. Furthermore, AFM-based SMFS has demonstrated 0.5-μs temporal resolution in an individual record (Rico, Gonzalez, Casuso, Puig-Vidal, & Scheuring, 2013), which is 10–50-fold faster than is achieved in advanced dual-beam optical traps (Neupane et al., 2016; Zoldak, Stigler, Pelz, Li, & Rief, 2013). Thus, there is significant opportunity to broaden the role of AFM in biophysics by complementing AFM’s existing strengths with improved force precision and stability, while maintaining (or even improving) its temporal resolution. AFM’s relatively poor force stability has been previously attributed to tipsample motion. In other words, the mechanical frame of the AFM was not stable enough. Recently, we showed force stability in a modern compact AFM (Cypher, Asylum Research) is not limited by tip-sample stability but rather by the mechanical properties of the AFM cantilever (Churnside et al., 2012). In particular, the primary source of force drift was the cantilever’s gold coating, even though it was coated on both sides to minimize thermally induced bimetallic artifacts. The key to improving short-term force precision, in contrast, has long been known: reduce the hydrodynamic drag of the cantilever (Viani et al., 1999). This result is a consequence of the fluctuation–dissipation theorem: ΔF¼√(4kBTβΔf ) where ΔF is the force precision, kBT is the thermal energy, β is the hydrodynamic drag of the cantilever, and Δf is the bandwidth of the measurement. Reducing β at constant cantilever stiffness (k) has the additional benefit of improving temporal resolution by increasing the speed at which the cantilever responds to a change in force (τ ¼ β/k in the overdamped regime). The standard approach to reducing β is to reduce the cantilever’s size. However, shorter cantilevers are inherently stiffer, and recent work from our lab has demonstrated that stiffer cantilevers suffer from decreased force stability (Bull et al., 2014). This poor force stability results from positional noise (Δx) in the optical lever arm, which results in larger instrumentation-induced force noise (ΔF ¼ kΔx) for stiff cantilevers (Sullan, Churnside, Nguyen, Bull, & Perkins, 2013). Thus, long, soft uncoated cantilevers are better suited for experiments that require

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excellent force stability, at the cost of decreased temporal resolution and force precision (Edwards & Perkins, 2016). Alternatively, experiments focused on measuring rapid or subtle conformational changes benefit from shorter, stiffer cantilevers at the cost of force stability. Hence, a careful choice and indeed a compromise is required when choosing among commercially available cantilevers. In other words, there is no commercially available cantilever that is best for all AFM-based SMFS applications. The challenge to achieving an optimal cantilever is that the cantilever needs to be both short and soft. Hence, one needs to circumvent the scaling relation for a rectangular cantilever’s stiffness: k∝wT 3 =L 3 , where w is the width of a cantilever, T is its thickness, and L is its length. As shown in Fig. 2, we bypassed this scaling relation by modifying the shape and thickness of the cantilevers using FIB milling (Bull et al., 2014), building upon earlier work (Hodges, Bussmann, & Hoh, 2001; Maali et al., 2006). An FIB is a common instrument on university campuses, often housed in shared nanofabrication facilities. Essentially, it allows one to sandblast a sample with gallium ions. Additionally, a scanning electron microscope (SEM) or FIB allows one to pattern the cantilever with a thin transparent material [tetraethyl orthosilicate (TEOS)]. We used this process to form a capping layer on top of the cantilever’s gold coating. This TEOS-based capping layer acted as a protective mask during a subsequent wet chemical etch used to remove the gold and underlying chromium, allowing us to preserve a small reflective patch at the free end of the cantilever. As a result, we retained the benefits of a gold-coated cantilever’s high reflectivity while avoiding the adverse effect of a gold coating on a cantilever’s force stability (Bull et al., 2014). An underappreciated concern in AFM-based SMFS is that many AFM cantilevers, particularly the shorter ones, are not overdamped, even when immersed in liquid and positioned 50 nm over a surface (Bull et al., 2014; Edwards et al., 2015). Yet, traditional SMFS analyses assume that force probes are overdamped (Bell, 1978; Dudko, Hummer, & Szabo, 2006; Evans & Ritchie, 1997; Merkel, Nassoy, Leung, Ritchie, & Evans, 1999). An underdamped cantilever “rings” due to its response to Brownian motion. Such oscillations mask small, subtle changes in structure and presumably affect folding and unfolding rates (Edwards et al., 2015; Edwards & Perkins, 2016) due to the exponential sensitivity of such rates to applied force. The performance gains of FIB modification are summarized in Fig. 3, which details the force power spectral density (PSD) (pN2/Hz) and force precision over a given averaging time (i.e., the Allan deviation) (Sullivan, Allan, Howe, & Walls, 1990). In particular, an FIB-modified BioLever Mini

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Fig. 2 A protocol for micromachined cantilevers. (A) Modification scheme for BioLever Minis. A frame for the gold mask is milled into the cantilever using an FIB. Next, the central region of the cantilever is removed. The thickness of the supports is reduced with the FIB. Then, a glass-like capping layer is deposited using electron-beam-induced deposition. Finally, a wet etch removes unprotected gold and chromium. (B) Modification scheme for BioLever Fasts. A small portion of the central region is removed. Next, the central region is further removed with cuts from the FIB to achieve the desired leg width. Finally, the thickness of the legs is reduced with the FIB to produce a straight cantilever.

cantilever (L ¼ 40 μm, k ¼ 7.5 pN/nm; Olympus) achieves sub-pN force noise to cover 5 decades of bandwidth without a substantial reduction in temporal resolution. Similarly, FIB modification of a BioLever Fast (L ¼ 10 μm, k ¼ 30 pN/nm, Olympus) maintained 1-μs-scale temporal

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Fig. 3 Comparing the mechanical properties of standard and FIB-modified cantilevers in liquid. (A) The force power spectral density (PSD) of cantilevers in liquid is plotted as a function of frequency. The data were taken at 50 nm over the surface. The BioLever Mini (green) and modified BioLever Mini (light blue) have fundamental frequencies near 5 and 30 kHz, respectively. The BioLever Fast (dark red) and modified BioLever Fast (tan) rolloff frequencies are at 500 and 200 kHz. (B) Force precision for each cantilever was rffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffi Effi 1D ðx i + 1  x i Þ2 , where x i is the mean calculated from the Allan deviation σ x ðT Þ ¼ T 2 value of the data over the ith time interval T. The Allan deviation represents the average force noise over a given averaging time derived from the same set of data used in (A). We note that, at the very shortest times, the motion of the cantilever becomes correlated, distorting the Allan deviation. This region of the curve is deemphasized using a dotted line.

resolution and improved its force stability, while achieving an overdamped cantilever. Hence, the modification process described here is a straightforward way to optimize AFM performance.

2. OVERVIEW OF MODIFICATION PROCESS Our protocol has been optimized to improve throughput and simplicity. We note that once trained, undergraduates in the lab routinely modify

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2–4 cantilevers per hour, depending on the cantilever size. Such modifications do require expertize in operating an FIB. For those with prior FIB experience, proficiency in implementing this protocol can be achieved in 10–20 h of FIB time. For those without prior FIB experience, an additional 10–24 h of training should be expected. The modification process for the BioLever Mini is described in Fig. 2A. Initially, a thin rectangular-shaped frame is milled through the gold at the free end of the cantilever to prevent undercutting of the transparent-capping layer during a subsequent wet chemical etch (Fig. 4). Next, the central portion of the cantilever is removed, and the remaining supports are thinned. This pair of steps reduces the hydrodynamic drag and stiffness of the cantilever. The thinning step also reduces stress-induced cantilever bending and is used to straighten the cantilever. Next, an electron-beam-induced transparent-capping mask is patterned using the FIB’s gas injection system (GIS) and SEM. Finally, the cantilever’s gold and underlying chromium layers are chemically etched with the transparent-capping layer to preserve a small reflective patch at the cantilever’s free end. The modification process for the BioLever Fast is described in Fig. 2B. Removal of the center portion is conducted in two steps: (i) removing a small rectangular portion and (ii) widening that portion by

Fig. 4 Improving performance of TEOS capping layer by milling a trench into the gold coating. (A) The trench creates a frame for the TEOS to adhere to the underlying silicon nitride, thus preventing undercutting of the gold. (B) An incomplete or nonexistent trench may allow the etchant to remove gold underneath the TEOS patch and can cause undercutting and a reduction or loss of the gold reflective patch. (C) A complete trench will be milled down to silicon nitride and will have no gold remaining within the 500-nm thickness of the trench. (D) An incomplete trench mill will have areas of gold visible, and will prevent maximum protection. (E) A full 500-nm thick trench surrounding a 12  8 μm2 area.

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narrowing the supports. This process avoids stress-induced buckling that can occur when cutting out a larger section of ultrashort cantilevers in one step. As with the BioLever Mini, thinning of the supports is used to reduce the stiffness and straighten cantilevers. For these ultrashort cantilevers, the major challenge during modification is minimizing ion-beam-induced bending of the cantilever. We found that even a single image acquired using the FIB (but not the SEM) can induce noticeable bending.

3. METHODS AND PROTOCOLS 3.1 Equipment and Materials 1. Dual-beam SEM/FIB (Anova 600 Nanolab, FEI). 2. GIS for FIB including tetraethyl orthosilicate (TEOS) (GIS, FEI). 3. Atomic force microscope (Cypher, Asylum Research). a. A modified BioLever Fast will require an ultrasmall detection laser (ours is 3  3 μm2) (Edwards et al., 2015). We successfully detect the modified BioLever Mini with laser spots of 30  10 μm2, 10  3 μm2, and 3  3 μm2. 4. Microscope with 10–50 objectives (BH2-UMA with NeoSPlan objectives, Olympus). 5. Stereo microscope (Vistavision, VWR). 6. AFM cantilevers (AC10DS BioLever Fasts and AC40TS BioLever Minis, Olympus). 7. 12.7-mm diameter, low-profile 90-degree SEM pinstubs (16171, Ted Pella). 8. Pinstub boxes (16140 (holds 4), 16630 (holds 1), Ted Pella). 9. Carbon tape (16084-7, Ted Pella). 10. SEM mount gripper (1664, Ted Pella). 11. Paddle-style tweezers (SM109SA, Techni-Tool). 12. Straight tweezers (3C-SA-PI or 2A-SA-SE, Excelta). 13. Gold etchant (gold Etchant TFA, Transene). 14. Chromium etchant (chrome Etchant 1020, Transene). 15. 99.9% isopropyl alcohol (IPA) (A416-4, Fisher). 16. Ultrapure water (>18 MΩ, Millipore). 17. 30% Hydrogen peroxide (5240-05, Macron). 18. Nitrile gloves (1100, Handpro). 19. 12-mm mica surface (50-12, Ted Pella) mounted on specimen discs (16218, Ted Pella).

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3.2 Mounting and Loading AFM Cantilevers 3.2.1 Mounting AFM Cantilevers on Pinstub 1. Always wear gloves when handling anything placed in the FIB to prevent oil and dust from contaminating the chamber. 2. Secure a 90-degree pinstub in a pinstub box. 3. Place a small (10 mm  3 mm) piece of double-sided carbon tape at the edge of the pinstub (Fig. 5). a. Carbon tape is conductive and therefore electrically grounds the cantilever to the pinstub. b. Carbon tape can be reused until it no longer holds the cantilevers securely. 4. Using tweezers, place the cantilever chip inverted (top-side facing down) on the carbon tape so that about 1/2 of the chip is on the carbon tape with the cantilever over the overhang. We use straight tweezers to grasp the chip from top and bottom away from the tip (Fig. 5). 5. Press down on the back of the chip gently with tweezers to level the chip on the stub.

Fig. 5 Mounting cantilevers for FIB modification. Image shows a single SEM pinstub loaded with three AFM chips. Chips are placed upside down onto carbon tape using tweezers. With the chip upside down, the tip of the cantilever is downward to ensure milled material is not redeposited on the tip. The cantilever position on the carbon tape is staggered to avoid chips blocking access to neighboring cantilevers. The inset shows a cartoon of typical positioning of the cantilevers.

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6. Load up to five chips on this stub. Each chip should be placed further on the carbon tape than the preceding chip (Fig. 5). a. Aligning the chips in this way ensures access to one cantilever is not blocked by another chip. 3.2.2 Mounting Pinstub in FIB 1. Vent the FIB to bring the chamber to atmospheric pressure. 2. Load the pinstubs into the FIB so that the chips are oriented perpendicular to the plane defined by the SEM and FIB columns (Fig. 6). a. In this orientation when the stage is rotated to place the FIB at normal incidence, the SEM can clearly visualize cantilever bending. 3. Ensure the pinstubs are secured tightly in the holder. 4. Reseal the chamber and initiate pump-down.

3.3 Imaging with the SEM and Preparing the FIB To begin, we describe using SEM imaging to orient and position the AFM cantilevers properly for concurrent SEM imaging and FIB milling. We then overlap the SEM and FIB fields of view. A summary of FIB-/SEMoperating parameters is given in Table 1.

Fig. 6 Orientation of the cantilevers in the FIB/SEM machine. The electron beam and the ion beam are coplanar in the y–z plane. The angle between the two beams is 52 degrees. The stage holding the cantilevers rotates about the x-axis to modify the incidence angle of the beams on the cantilevers. The inset shows that the cantilevers are oriented along the x-axis. With this orientation the full length of the cantilever remains visible when rotated to normal incidence with the FIB beam for milling.

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Table 1 Standard Parameters for FIB Modification of Cantilevers. SEM FIB B.L. Mini Imaging SEM TEOS FIB Search Milling

FIB B.L. Fast Milling

Voltage (keV) 5

5

30

30

30

Current (pA)

400

400

1.5

93

9.7

Dwell time (ns/pixel)

300

200

300, 1000

200

200

HFW (μm)

Variable

25

Variable

25–45

10–15

Pixels

1024  884 1024  884 1024  884 1024  884

1024  884

Overlap

N/A

0%

N/A

50%

50%

Interaction diameter

N/A

50%

N/A

0%

0%

Imaging averages

User N/A preference

Single frame

Single frame Single frame

Depth

N/A

N/A

5 μm

5 μm

5 μm

3.3.1 Imaging with the SEM 1. Standard SEM imaging parameters are 5 kV, 400 pA, 300 ns/pixel, 1  frame averaging, 1024  884 pixels (Table 1). We use the EverhartThornley detector for all imaging, except to check contrast of the TEOS patch. 2. Zero the SEM beam offset. 3. Begin imaging and move stage to center a cantilever in the field of view. 4. Locate a cantilever with the SEM and optimize imaging with a horizontal field width (HFW) less than 100 μm. 5. If necessary, link the z-stage height to the focused working distance to calibrate the z-stage properly. 6. Physically rotate the stage to the correct orientation (Fig. 6). 3.3.2 Finding Eucentric Height and Coincident Point We set the stage height so that the SEM and FIB beams irradiate the same location (coincidence position), and so that when the sample is tilted, the specimen-aperture distance does not change (eucentric height). In our dualbeam SEM/FIB (Anova 600 Nanolab, FEI), the coincident position and eucentric height are the same location. Achieving the correct eucentric

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and coincident height allows concurrent SEM imaging and FIB milling, and properly positions the needle when using the GIS. 1. Safely move the stage to a height near the expected eucentric height. In our case, this is a working distance of 5 mm. 2. Focus on the specimen at a stage tilt of 0 degrees with a 100 μm HFW. 3. Place crosshairs on a visible feature such as the long edge of the cantilever. 4. Tilt the specimen to 10 degrees. 5. By adjusting only the stage height (not the x-, y-stages) move the feature back under the crosshairs. 6. Tilt the specimen to the angle such that the stage is perpendicular to the FIB. This angle is 52 degrees for our SEM/FIB. 7. Adjust the stage height until the feature is back under the crosshairs. Specimen is now at eucentric and coincident height. 3.3.3 Finding the Cantilever with the FIB For concurrent SEM imaging and FIB milling, we overlap the SEM and FIB beams by shifting the SEM beam. The challenge is to locate the cantilever using the FIB without significantly dosing the cantilever, which will induce bending. 1. Center the cantilever in the SEM while tilted to 52 degrees. 2. Zero the FIB offset. 3. Set FIB to lowest magnification (HVW: 640 μm) and set standard FIB settings for finding the cantilever: 1.5 pA, 30 kV, 300 ns/pixel, 1024  884 pixels (Table 1). The dwell time can be increased to 1 μs to improve imaging if necessary. 4. Set averaging to integrate one frame so that a single image is taken at a time, stopping after each. a. This setting is important as it minimizes dosing of the cantilever with ions by preventing multiple, repeated images from being acquired. 5. Use a reduced-scan area (RSA) to image a small portion of the AFM chip far from the cantilever at the top or bottom fifth of the screen. 6. Adjust the focus of the FIB beam. a. Focus can be tuned at higher magnification, so long as the RSA is used to ensure the cantilever is not imaged. 7. To achieve the lowest possible dosage, locate the cantilever by taking a single image with the FIB at the lowest magnification (in our machine, HVW: 640 μm).

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8. The cantilever should be visible at the center of the chip edge. Center the stage on the cantilever. a. At this low current and magnification, the dosing of the cantilever is minimal. b. If the cantilever is difficult to see, another technique for finding the cantilever with minimal dosing uses the RSA to search just along the edge of the chip at higher magnification. 9. Zoom in to 25 μm HFW for BioLever Fasts or 100 μm HFW for the BioLever Mini. 10. Use the RSA to find the very base of the cantilever along the edge of the chip without dosing the cantilever itself. a. By imaging along the chip, we only apply a dose at the very base of the cantilever. Though imaging the base of the lever can induce bending, this bending is generally eliminated by a subsequent thinning process. 11. Identify the wings of the cantilever from the image. For the BioLever Mini, these wings are clearly visible on either side of the cantilever (Fig. 7). For the BioLever Fast, there is a slight protrusion at the base of the cantilever. If wings are not visible, slowly expand the RSA beyond the chip to observe them. 12. Center on a wing and zoom in so only the wing is visible (Mini: 8 μm HFW, Fast: 2 μm HFW). 13. Set ion-beam current to the desired current: 93 pA for BioLever Mini, 9.7 pA for BioLever Fast. a. In our hands, when the current is changed, the wing remains visible or can be relocated easily. If necessary, we refer you to your protocols for aligning the FIB using different apertures. b. It is important to check the current output to assure correct milling times. Beam currents may change from day to day and evolve over the lifetime of the apertures. If in doubt, use a Faraday cup to determine the current. 14. Take a single FIB image of the wing and recenter if necessary. 15. Optimize FIB parameters with individual images. a. Remember that the FIB is inherently destructive and will actively change the shape of the wing as it is imaged. If possible, start on a small portion of the wing and, when the focus is presumed to be correct, move to another less-imaged portion of the wing to check. b. We utilize these wings for optimizing the FIB focus without dosing the cantilever. While the focus can be optimized elsewhere, the benefit is that these wings are at the same vertical height as the cantilever itself.

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Fig. 7 Modification of a BioLever Mini cantilever using an FIB. SEM images taken at each step of the FIB process and after gold removal. All images were taken at a 52-degree perspective, with the cantilevers normal to the ion beam during modification. (A) SEM image of an unmodified BioLever Mini. (B) Image after the TEOS frame was milled into the cantilever. (C) Image after a rectangular pattern was cut to define the area to be removed. (D) Image after a cut along the chip caused the flap to fold up. (E) Image after the supports were thinned to reduce cantilever bending and stiffness. (F) Image after the metal etch removed the gold and chromium (fully modified cantilever). The dark rectangle is indicative of material contrast and shows the preservation of the gold patch after etching.

16. Zoom out to 50 μm HFW for BioLever Mini, 15 μm HFW BioLever Fast, and use RSA to find and center on the cantilever as in step 8. 17. With the SEM, image the cantilever and use x-, y-beamshifts to center image. Do not move the stage, or you will have to recenter with the FIB.

3.4 Modifying BioLever Minis with the FIB The cantilever is milled and thinned to reduce both stiffness and hydrodynamic drag, and a protective TEOS layer is added to preserve a gold patch at the end of the cantilever. 3.4.1 Creating TEOS Frame In order to improve adhesion of the TEOS capping layer, we remove the gold immediately surrounding the TEOS patch (Figs. 4 and 7B). We find this improves the robustness of the TEOS patch (Fig. 4A and B), presumably by preventing undercutting of the gold.

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1. Use the RSA to image the cantilever out to its end. a. Ideally, this is the only process where the entire cantilever is imaged with the FIB. 2. Use FIB patterning tools to draw a 13  10 μm2 unfilled rectangle. 3. Place the rectangle at the center of the free end of the cantilever (Figs. 4E and 7B). 4. FIB parameters for cutting silicon are typically suitable for milling silicon nitride, though the conversion from dose to depth will differ. For BioLever Minis, we use 30 keV, 93 pA, 200 ns/pixel dwell time, 1024  884 pixels, 50% beam overlap (Table 1). We select a depth of 5 μm, which is much larger than the thickness of the cantilever. Rather than calibrate the precise depth, we set the depth to much deeper than necessary and manually control the time by observing in the SEM. 5. Use blur or defocus to achieve a 500 nm wide beam. We use a defocus of 175 μm, but suggest calibrating this on different SEM/FIBs. a. The beam width can be empirically measured by cutting into the wings. 6. Turn on the SEM and center and magnify the image on the end of the cantilever (20 μm HFW). a. SEM imaging during milling allows optimal control of timing as well as providing warning of problems. 7. To ensure a rapidly updated SEM image, use a dwell time of 100 ns/ pixel without averaging. 8. Before patterning, take a single FIB image and realign the pattern if necessary. 9. Begin SEM imaging and start FIB patterning. 10. Stop pattern when SEM shows the gold is fully removed. a. Look for when the final beads of gold disappear to ensure the gold is milled completely without milling too far into the silicon nitride of the cantilever (Fig. 4C vs D).

3.4.2 Creating the Legs of the Cantilever To reduce hydrodynamic drag and stiffness, we remove a large central portion of the cantilever by cutting out three sides and then allowing it to fold up as a flap. 1. Using the RSA, take a single FIB image from the chip out to the beginning of the TEOS frame. This covers the entire flap area.

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2. Draw an unfilled rectangle pattern extending 1 μm from edge of the frame and 1 μm onto the chip. Use the measurement tools to leave 1-μm legs (Fig. 7C). Ensure only the perimeter will be milled. a. Preserving the frame requires that the flap must be milled at least several hundred nm from the frame. b. This pattern is drawn on the chip so that one side of the rectangular cut will be incomplete. 3. Use our standard milling parameters for a BioLever Mini (Table 1). 4. Defocus should be set to 0 to achieve the smallest possible beam diameter. 5. Ensure the position of the rectangle has not changed by taking another RSA image. 6. Activate and optimize SEM imaging of the area being milled. 7. Begin patterning, and stop when the flap separates from the cantilever. 8. Take a single FIB image and check that the flap is detached on three sides. a. If detachment is incomplete before continuing, there is a high risk that the flap will fail to release. 9. If the flap is not detached, you may continue the patterning. However, a faster approach is to replace the rectangular pattern with single lines patterned across the attached region. Ensure multiple patterns are milled in parallel. 10. Stop pattern when flap is detached on three side. 11. Delete all patterns. 12. Draw a single line along the edge of the chip connecting the two cut lines. 13. Start SEM imaging and then start pattern. Stop pattern when the flap lifts, typically