Biomotors and their Nanobiotechnology Applications 9780367196134, 9781032313146, 9780429203367

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Biomotors and their ­Nanobiotechnology Applications This book – a collection of reviews and research articles by the top academics in the field – provides a glimpse of the cutting-edge technology and research being carried out and shows how researchers are utilizing this knowledge to develop new areas of study and novel applications. It serves as a valuable resource while exploring the latest advances in virus particle assembly and demonstrating how the knowledge of fundamental processes has been used to advance bio-nanotechnology. Chapters detail biophysical approaches and biomotor research, discusses the latest advances in DNA/RNA nanoparticle assembly and use, and introduces the use of DNA/RNA nanoparticles for drug delivery.

Biomotors and their ­Nanobiotechnology Applications

Edited by

Peixuan Guo and Aibing Wang

First edition published 2024 by CRC Press 2385 NW Executive Center Drive, Suite 320, Boca Raton FL 33431 and by CRC Press 4 Park Square, Milton Park, Abingdon, Oxon, OX14 4RN CRC Press is an imprint of Taylor & Francis Group, LLC © 2024 selection and editorial matter, Peixuan Guo and Aibing Wang; individual chapters, the contributors Reasonable efforts have been made to publish reliable data and information, but the author and publisher cannot ­ ublishers assume responsibility for the validity of all materials or the consequences of their use. The authors and p have attempted to trace the copyright holders of all material reproduced in this publication and apologize to ­copyright ­holders if permission to publish in this form has not been obtained. If any copyright material has not been ­acknowledged please write and let us know so we may rectify in any future reprint. Except as permitted under U.S. Copyright Law, no part of this book may be reprinted, reproduced, transmitted, or utilized in any form by any electronic, mechanical, or other means, now known or hereafter invented, ­including ­photocopying, microfilming, and recording, or in any information storage or retrieval system, without written ­permission from the publishers. For permission to photocopy or use material electronically from this work, access www.copyright.com or contact the Copyright Clearance Center, Inc. (CCC), 222 Rosewood Drive, Danvers, MA 01923, 978-750-8400. For works that are not available on CCC please contact [email protected] Trademark notice: Product or corporate names may be trademarks or registered trademarks and are used only for identification and explanation without intent to infringe. ISBN: 978-0-367-19613-4 (hbk) ISBN: 978-1-032-31314-6 (pbk) ISBN: 978-0-429-20336-7 (ebk) DOI: 10.1201/9780429203367 Typeset in Times by codeMantra

Dedicated to Steven Qian.

Contents Preface...............................................................................................................................................xi Authors ........................................................................................................................................... xiii Contributors ..................................................................................................................................... xv Note to Reader ................................................................................................................................xxi

PART I

Fundamental Mechanism of Biomotor Action

Chapter 1

Biological Nanomotors with Linear, Rotation, or Revolution Motion Mechanism .....3 Peixuan Guo, Hiroyuki Noji, Christopher M. Yengo, Zhengyi Zhao, and Ian Grainge

Chapter 2

Classifications and Typical Examples of Biomotors .................................................. 43 Peixuan Guo and Zhengyi Zhao

Chapter 3

Structure of Revolving Biomotors .............................................................................. 53 Peixuan Guo and Zhengyi Zhao

Chapter 4

Structure of Rotation Motors ..................................................................................... 59 Peixuan Guo and Zhengyi Zhao

Chapter 5

Structure of Linear Motors......................................................................................... 63 Peixuan Guo and Zhengyi Zhao

Chapter 6

Mechanical Properties of Molecular Motors and the Relevance to Their Biological Function........................................................................................... 67 Yuchuan Zheng and Jingyuan Li

Chapter 7

Molecular Mechanism of AAA-ATPase Motor in the 26S Proteasome .................... 81 Shuwen Zhang and Youdong Mao

Chapter 8

General Mechanism of Biomotors .............................................................................99 Peixuan Guo and Zhengyi Zhao

Chapter 9

Mechanism of Revolving Motors ............................................................................. 103 Peixuan Guo and Zhengyi Zhao

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Contents

Chapter 10 Mechanism of Rotary Motors .................................................................................. 109 Peixuan Guo and Zhengyi Zhao Chapter 11 Mechanism of Linear Motion................................................................................... 117 Peixuan Guo and Zhengyi Zhao Chapter 12 Finding of Widespread Viral and Bacterial Revolution dsDNA Translocation Motors Distinct from Rotation Motors by Channel Chirality and Size ................... 123 Gian Marco De-Donatis, Zhengyi Zhao, Shaoying Wang, Lisa P. Huang, Chad Schwartz, Oleg V. Tsodikov, Hui Zhang, Farzin Haque, and Peixuan Guo Chapter 13 The ATPase of the phi29 DNA Packaging Motor Is a Member of the Hexameric AAA+ Superfamily ............................................................................... 141 Chad Schwartz, Gian Marco De Donatis, Huaming Fang, and Peixuan Guo Chapter 14 Arginine Finger Serving as the Starter of Viral DNA Packaging Motors ............... 155 Chenxi Liang, Chun Chan, Zhefeng Li, Xiaolin Cheng, and Peixuan Guo Chapter 15 Three-Step Channel Conformational Changes Common to DNA Translocases of Bacterial Viruses T3, T4, SPP1, and phi29 .................................... 159 Shaoying Wang, Zhouxiang Ji, Erfu Yan, Farzin Haque, and Peixuan Guo Chapter 16 Sequence Dependence of Reversible CENP-A Nucleosome Translocation ............. 173 Micah P. Stumme-Diers, Thomas Stormberg, and Yuri L. Lyubchenko Chapter 17 Same Function from Different Structures among pac Site Bacteriophage (TerS) Terminase Small Subunits........................................................................................ 179 Lindsay W. Black and Krishanu Ray Chapter 18 Kinetic Study of the Fidelity of DNA Replication with Higher-Order Terminal Effects ....................................................................................................... 183 Yao-Gen Shu Chapter 19 Multilevel Control of the Activity of p97/Cdc48, A Versatile Protein Segregase ...207 Di Xia and Wai Kwan Tang Chapter 20 High-Resolution Structure of Hexameric Herpesvirus DNA Packaging Motor Elucidates Revolving Mechanism and Ends 20-Year Fervent Debate ..................... 227 Peixuan Guo

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Contents

PART II

Methods for the Study of Biomotors

Chapter 21 Methods for Single-Molecule Sensing and Detection Using Bacteriophage Phi29 DNA Packaging Motor .................................................................................. 235 Farzin Haque, Hui Zhang, Shaoying Wang, Chun-Li Chang, Cagri Savran, and Peixuan Guo Chapter 22 Instrumental Design for Five-Dimensional Single-Particle Rotational Tracking .... 257 Kuangcai Chen, Xiaodong Cheng, and Ning Fang Chapter 23 The Appropriate Ratio of Retroviral Structural Proteins Is Activated by the Spleen Necrosis Virus Post-Transcriptional Control Element ........................... 265 Gatikrushna Singh, Deepali Singh, Nicole Placek, Stacey Hull, Radhakrishna Sura, and Kathleen Boris-Lawrie

PART III Application of Biomotors Chapter 24 Translation of the Long-Term Fundamental Studies on Viral DNA Packaging Motors into Nanotechnology and Nanomedicine .................................................... 281 Chenxi Liang, Tao Weitao, Lixia Zhou, and Peixuan Guo Chapter 25 Translocation of Peptides through Membrane-Embedded SPP1 Motor Protein Nanopores .................................................................................................... 321 Shaoying Wang, Farzin Haque, and Peixuan Guo Chapter 26 Insertion of Channel of phi29 DNA Packaging Motor into Polymer Membrane for High-Throughput Sensing ................................................................ 333 Zhouxiang Ji and Peixuan Guo Chapter 27 Engineering of Protein Nanopores for Sequencing, Chemical or Protein Sensing, and Disease Diagnosis .................................................................. 343 Shaoying Wang, Zhengyi Zhao, Farzin Haque, and Peixuan Guo Chapter 28 Phage Portal Channels as Nanopore Sensors ........................................................... 357 Farzin Haque and Shaoying Wang Chapter 29 Controlled Co-assembly of Viral Nanoparticles of Simian Virus 40 with Inorganic Nanoparticles: Strategies and Applications ............................................. 367 Wenjing Zhang, Xian-En Zhang, and Feng Li

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Chapter 30 Potential of 3Dpol as an Enzymatic Reader for Direct RNA Sequencing ............... 375 Howard B. Gamper, Sitao Yin, and Ya-Ming Hou Chapter 31 Channel from Bacterial Virus T7 DNA Packaging Motor for the Differentiation of Peptides Composed of a Mixture of Acidic and Basic Amino Acids................................................................................................... 383 Zhouxiang Ji and Peixuan Guo Chapter 32 Nano-channels of Viral DNA Packaging Motor as Single Pore to Differentiate Peptides with Single-Amino Acid Difference .......................................................... 395 Zhouxiang Ji, Xinqi Kang, Shaoying Wang, and Peixuan Guo Index ..............................................................................................................................................407

Preface Biomotors, also known as biological motors, have become an advanced topic in both fundamental research and application practice. Biomotors are a group of protein ATPases that convert chemical energy to mechanical motion in all kinds of motion activities (such as breathing, walking, speaking, hearing, blinking, and heartbeat) and in other critical biological processes (such as cell mitosis, binomial fission, DNA replication, RNA transcription, cargo delivery, cellular transport, macromolecule translocation, viral infection, and flagella motion). Studies about the mechanisms and functions of these biomotors are essential to not only answer the question that how life works but also develop new treatments and therapeutic targets towards different kinds of diseases. However, currently, the mechanism of many motors remains unknown. With the development of new technologies and decades of hard work, the field of biomotors is entering a new era. New methods such as categorized Cryo electron microscopy (cryo-EM) and single-molecule imaging offer us a much clearer perception of the structure of different kinds of motors, for example, MCM helicase, V1-ATPase, and ClpX. Based on these studies, numerous hypotheses are proved, while growing new ideas are generated. Recently, a new revolution motion mechanism, other than the traditionally linear and rotation motion process, has been reported. Revolving motors have been found to apply to a wide range of ATPase motors in the translocation of lengthy genomic dsDNA of prokaryotes such as Ftsk in Gram-negative bacteria, SpoIIIE in Grampositive bacteria, plasmid conjugation ATPase TraB in Streptomyces, dsDNA packaging motors in bacteriophages, as well as of eukaryotes such as the dsDNA packaging motors in herpesviruses, poxvirus, and adenovirus, and mimivirus. It is expected that the chromosomal dsDNA translocation and processing ATPases use the same removing mechanism since it is not realistic to use the rotation mechanism to transport the lengthy human genome without coiling and tangling. If the rotation mechanism had been used for the transporting of the human dsDNA genome, huge energy would have been required to resolve the supercoil. Nature has evolved an elegant and novel revolving mechanism without coiling. The use of the asymmetrical ATPase hexamers to process their substrates as reported in phi29 dsDNA packaging motors has been reported recently to be common for many ATPases in prokaryotes, eukaryotes including human. The booming new literature about biomotor studies would shine new lights and add a new color to this field and promote a wide field of applications. Due to the significant advancement in biomotor studies, it is time to translate the original research into a textbook. We hope this book can serve as a gate-pointer for senior students, graduate school attendees, postdoctoral researchers, academic faculties, and industry scientists from all kinds of other sciences to expand their knowledge and to advance the exciting and critical yet not well-developed field. It is imperative that we work together to propel the biomotor studies towards the target or beyond. We hope readers will find this good resourceful. Peixuan Guo Fellow of the National Academy of Inventors (NIA) The Ohio State University, Columbus, OH Spring 2021

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Authors Peixuan Guo, a pioneer of RNA nanotechnology, has held three endowed chair positions at three different prestigious universities. Currently, he is the Sylvan G. Frank Endowed Chair in Pharmaceutics and Drug Delivery and the Director of the Center for RNA Nanobiotechnology and Nanomedicine at The Ohio State University (OSU). He is the President of the International Society of RNA Nanotech and Nanomedicine (myisrnn.org). He received his Ph.D. from the University of Minnesota in 1987 and conducted his postdoctoral training at NIH under Bernard Moss (a member of the US NAS and the pioneer of vaccinia virus as a vaccine vector). He joined Purdue University in 1990, tenured in 1993 and became a full Professor in 1997, and honored as a Purdue Distinguished Faculty Scholar in 1998. He served as the Director of the NIH Nanomedicine Develoment Center (NDC) from 2006 to 2011. Also, he was the Director of NCI Cancer Nanotech Platform Partnership Program from 2012 to 2017. To date, he has invented 70 patents (13 granted and 57 in Provisional and PCT), 252 high-impact publications, with an H-index of 71 and cited 16,905 on Google Scholar and has an H-index of 57 and cited 10,765 on Web of Science. He has been  extensively involved in the research of DNA packaging motors, biomotors, and nanomachines for many years since he elucidated the phi29 packaging motor components, interactions, and proposed mechanism of packaging. His achievements in Biomotors led to his award as the 2021 Innovator of the Year at The Ohio State University and his election as an NAI (National Academy of Inventors) Fellow in 2022. Aibing Wang received his Bachelor’s and Master’s degrees from Hunan Agricultural University and obtained his Ph.D. in Biochemistry & Molecular Biology from the Chinese Academy of Medical Sciences (CAMS) & Peking Union Medical College (PUMC) in 2005. Thereafter, Dr Wang joined in National Heart, Lung and Blood Institute (NHLBI), National Institutes of Health (NIH) and worked there as Visiting Fellow, Research Fellow, and Biologist for over 10 years. In the following years, he has been a Senior Scientist at PCB Biotechnology, LLC. and a distinguished Professor of “Shenong Scholar program” and “Furong Scholar Program” at Hunan Agricultural University. Dr Wang has wide research interests ranging from the mechanisms or gene functions associated with virus infection & vaccine development, molecular motors, cardiovascular & metabolic disorders, oxidative stress, gene editing, antiviral chemicals & food toxins, to tumorigenesis and targeted delivery of therapeutics, etc., using animal models, if required, established by embryonic stem cell (ES)-based gene targeting or CRISPR/Cas9-mediated technology. His outstanding achievements include A) identification of so far the highest gene targeting efficiency locus (>95% vs 1–10% in most cases) and application of it for targeted transgenesis; B) design and generation of a series of genetically modified mouse models (>10) using traditional ES cells based HR mediated or CRISPR/ Cas9 mediated approach; C) expression of full-length nonmuscle myosin II macromolecular protein complex (>500kDa) using baculovirus system in the field; D) development of effective detection methods and vaccines for animal viruses such as canine papillomaviruses (CPVs). So far, he has published over 70 scientific papers, invented three issued patents, acquired three grants and obtained one Provincial Award. Dr Wang has also served as a reviewer for 20 journals.

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Contributors Lindsay W. Black Department of Biochemistry and Molecular Biology University of Maryland School of Medicine Baltimore, Maryland Kathleen Boris-Lawrie Department of Veterinary and Biomedical Sciences, College of Veterinary Medicine University of Minnesota Saint Paul, Minnesota and Department of Veterinary Biosciences, Center for Retrovirus Research The Ohio State University Columbus, Ohio Chun Chan Center for RNA Nanobiotechnology and Nanomedicine College of Pharmacy and College of Medicine James Comprehensive Cancer Center Dorothy M. Davis Heart and Lung Research Institute College of Pharmacy, Biophysics Graduate Program, Translational Data Analytics Institute The Ohio State University Columbus, Ohio Chun-Li Chang Birck Nanotechnology Center School of Mechanical Engineering Weldon School of Biomedical Engineering Purdue University West Lafayette, Indiana Kuangcai Chen Imaging Core Facility and Department of Chemistry Georgia State University Atlanta, Georgia

Xiaolin Cheng Center for RNA Nanobiotechnology and Nanomedicine College of Pharmacy and College of Medicine James Comprehensive Cancer Center Dorothy M. Davis Heart and Lung Research Institute College of Pharmacy, Biophysics Graduate Program, Translational Data Analytics Institute The Ohio State University Columbus, Ohio Gian Marco De-Donatis Nanobiotechnology Center Department of Pharmaceutical Sciences, College of Pharmacy Markey Cancer Center University of Kentucky Lexington, Kentucky Dana Driver Center for RNA Nanobiotechnology and Nanomedicine College of Pharmacy and College of Medicine Dorothy M. Davis Heart and Lung Research Institute Comprehensive Cancer Center The Ohio State University Columbus, Ohio Huaming Fang Nanobiotechnology Center College of Pharmacy Markey Cancer Center University of Kentucky Lexington, Kentucky Ning Fang Xiamen University Xiamen, China

Xiaodong Cheng Wenzhou Medical University Wenzhou, China

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Howard B. Gamper Department of Biochemistry and Molecular Biology Thomas Jefferson University Philadelphia, Pennsylvania Ian Grainge School of Environmental and Life Sciences University of Newcastle Callaghan, Australia Peixuan Guo Nanobiotechnology Center Markey Cancer Center Department of Pharmaceutical Sciences, College of Pharmacy William Farish Endowed Chair in Nanobiotechnology, School of Pharmacy University of Kentucky Lexington, Kentucky and Center for RNA Nanobiotechnology and Nanomedicine Division of Pharmaceutics and Pharmaceutical Chemistry, College of Pharmacy Department of Physiology & Cell Biology, College of Medicine James Comprehensive Cancer Center Dorothy M. Davis Heart and Lung Research Institute Biomedical Science Graduate Program, College of Medicine The Ohio State University Columbus, Ohio and Department of Biomedical Engineering College of Engineering and College of Medicine University of Cincinnati Cincinnati, Ohio

Contributors

Farzin Haque Division of Pharmaceutics and Pharmaceutical Chemistry, College of Pharmacy Department of Physiology & Cell Biology, College of Medicine Dorothy M. Davis Heart and Lung Research Institute The Ohio State University Columbus, Ohio and Nanobiotechnology Center Markey Cancer Center Department of Pharmaceutical Sciences, College of Pharmacy University of Kentucky Lexington, Kentucky and Department of Biomedical Engineering College of Engineering and College of Medicine University of Cincinnati Cincinnati, Ohio and P&Z Biological Technology Newark, New Jersey Ya-Ming Hou Department of Biochemistry and Molecular Biology Thomas Jefferson University Philadelphia, Pennsylvania Lisa P. Huang Institute for Biomarker Research Medical Diagnostic Laboratories, L.L.C. Hamilton, New Jersey Stacey Hull Department of Veterinary Biosciences, Center for Retrovirus Research The Ohio State University Columbus, Ohio

xvii

Contributors

Zhouxiang Ji Center for RNA Nanobiotechnology and Nanomedicine Division of Pharmaceutics and Pharmaceutical Chemistry, College of Pharmacy Department of Physiology & Cell Biology, College of Medicine Dorothy M. Davis Heart and Lung Research Institute James Comprehensive Cancer Center The Ohio State University Columbus, Ohio Peng Jing Department of Biomedical Engineering, College of Engineering and College of Medicine University of Cincinnati Cincinnati, Ohio Xinqi Kang Center for RNA Nanobiotechnology and Nanomedicine Division of Pharmaceutics and Pharmaceutical Chemistry, College of Pharmacy Department of Physiology & Cell Biology, College of Medicine Dorothy M. Davis Heart and Lung Research Institute James Comprehensive Cancer Center The Ohio State University Columbus, Ohio Feng Li State Key Laboratory of Virology, Wuhan Institute of Virology, Center for Biosafety Mega-Science Chinese Academy of Sciences Wuhan, China Jingyuan Li Zhejiang Province Key Laboratory of Quantum Technology and Device, Department of Physics, Institute of Quantitative Biology Zhejiang University Hangzhou, China

Zhefeng Li Center for RNA Nanobiotechnology and Nanomedicine College of Pharmacy and College of Medicine James Comprehensive Cancer Center Dorothy M. Davis Heart and Lung Research Institute The Ohio State University Columbus, Ohio Chenxi Liang Center for RNA Nanobiotechnology and Nanomedicine College of Pharmacy and College of Medicine James Comprehensive Cancer Center Dorothy M. Davis Heart and Lung Research Institute Biomedical Science Graduate Program, College of Medicine The Ohio State University Columbus, Ohio Yuri L. Lyubchenko Department of Pharmaceutical Sciences University of Nebraska Medical Center Omaha, Nebraska Youdong Mao Center for Quantitative Biology, School of Physics Peking University Beijing, China and Dana-Farber Cancer Institute Harvard Medical School Boston, Massachusetts Carlo Montemagno Department of Biomedical Engineering College of Engineering and College of Medicine University of Cincinnati Cincinnati, Ohio Hiroyuki Noji Department of Applied Chemistry The University of Tokyo Bunkyo City, Tokyo, Japan

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Contributors

Nicole Placek Department of Veterinary Biosciences, Center for Retrovirus Research The Ohio State University Columbus, Ohio

Gatikrushna Singh Department of Veterinary and Biomedical Sciences, College of Veterinary Medicine University of Minnesota Saint Paul, Minnesota

Krishanu Ray Department of Biochemistry and Molecular Biology University of Maryland School of Medicine Baltimore, Maryland

Thomas Stormberg Department of Pharmaceutical Sciences University of Nebraska Medical Center Omaha, Nebraska

Cagri Savran Birck Nanotechnology Center School of Mechanical Engineering Weldon School of Biomedical Engineering Purdue University West Lafayette, Indiana Chad Schwartz Nanobiotechnology Center Department of Pharmaceutical Sciences, College of Pharmacy Markey Cancer Center University of Kentucky Lexington, Kentucky Yao-Gen Shu Wenzhou Institute University of Chinese Academy of Sciences Beijing, China Deepali Singh Department of Veterinary Biosciences, Center for Retrovirus Research The Ohio State University Columbus, Ohio and School of Biotechnology Gautam Buddha University Greater Noida, India

Micah P. Stumme-Diers Department of Pharmaceutical Sciences University of Nebraska Medical Center Omaha, Nebraska Radhakrishna Sura Department of Veterinary Biosciences, Center for Retrovirus Research The Ohio State University Columbus, Ohio Wai Kwan Tang Laboratory of Cell Biology, Center for Cancer Research National Cancer Institute, National Institutes of Health Bethesda, Maryland Oleg V. Tsodikov Department of Pharmaceutical Sciences, College of Pharmacy University of Kentucky Lexington, Kentucky Anne P. Vonderheide Department of Biomedical Engineering, College of Engineering and College of Medicine University of Cincinnati Cincinnati, Ohio

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Contributors

Shaoying Wang Nanobiotechnology Center Department of Pharmaceutical Sciences, College of Pharmacy Markey Cancer Center University of Kentucky Lexington, Kentucky and Center for RNA Nanobiotechnology and Nanomedicine Division of Pharmaceutics and Pharmaceutical Chemistry, College of Pharmacy Department of Physiology & Cell Biology, College of Medicine Dorothy M. Davis Heart and Lung Research Institute James Comprehensive Cancer Center The Ohio State University Columbus, Ohio and P&Z Biological Technology Newark, New Jersey Tao Weitao Biology Department, College of Science and Mathematics Southwest Baptist University Bolivar, Missouri Di Xia Laboratory of Cell Biology, Center for Cancer Research National Cancer Institute, National Institutes of Health Bethesda, Maryland Erfu Yan Nanobiotechnology Center Markey Cancer Center Department of Pharmaceutical Sciences, College of Pharmacy University of Kentucky Lexington, Kentucky

Christopher M. Yengo Department of Cellular and Molecular Physiology College of Medicine, Pennsylvania State University Hershey, Pennsylvania Sitao Yin Department of Biochemistry and Molecular Biology Thomas Jefferson University Philadelphia, Pennsylvania Hui Zhang Nanobiotechnology Center Department of Pharmaceutical Sciences, College of Pharmacy Markey Cancer Center University of Kentucky Lexington, Kentucky Shuwen Zhang Center for Quantitative Biology, School of Physics Peking University Beijing, China Wenjing Zhang State Key Laboratory of Virology, Wuhan Institute of Virology, Center for Biosafety Mega-Science Chinese Academy of Sciences Wuhan, China and University of Chinese Academy of Sciences Beijing, China Xian-En Zhang Faculty of Synthetic Biology, Shenzhen Institute of Advanced Technology, Institute of Biophysics Chinese Academy of Sciences Beijing, China

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Zhengyi Zhao Nanobiotechnology Center Markey Cancer Center Department of Pharmaceutical Sciences, College of Pharmacy University of Kentucky Lexington, Kentucky and Center for RNA Nanobiotechnology and Nanomedicine College of Pharmacy and College of Medicine Dorothy M. Davis Heart and Lung Research Institute James Comprehensive Cancer Center The Ohio State University Columbus, Ohio and Nanobio Delivery Pharmaceutical Co. Ltd. Columbus, Ohio Yuchuan Zheng Zhejiang Province Key Laboratory of Quantum Technology and Device, Department of Physics, Institute of Quantitative Biology Zhejiang University Hangzhou, China

Contributors

Zhen Zheng Center for RNA Nanobiotechnology and Nanomedicine College of Pharmacy and College of Medicine Dorothy M. Davis Heart and Lung Research Institute Comprehensive Cancer Center The Ohio State University Columbus, Ohio Lixia Zhou Center for RNA Nanobiotechnology and Nanomedicine College of Pharmacy and College of Medicine Dorothy M. Davis Heart and Lung Research Institute Comprehensive Cancer Center The Ohio State University Columbus, Ohio

Note to Reader The chapters of this book are based on the extensive lifetime research efforts and collaboration of Dr. Peixuan Guo. Several chapters have been taken directly from various journal articles Dr. Guo has contributed to throughout his career. De Donatis, Gian Marco & Zhao, Zhengyi & Wang, Shaoying & Huang, Lisa & Schwartz, Chad & Tsodikov, Oleg & Zhang, Hui & Haque, Farzin & Guo, Peixuan. Finding of widespread viral and bacterial revolution dsDNA translocation motors distinct from rotation motors by channel chirality and size. Cell & Bioscience 4 (2014): 30. doi: 10.1186/2045-3701-4-30. Guo, Peixuan. “High resolution structure of hexameric herpesvirus DNA-packaging motor elucidates revolving mechanism and ends 20-year fervent debate.” Protein & Cell 11(5) (2020): 311–315. doi: 10.1007/ s13238-020-00714-w. Guo, Peixuan, Hiroyuki Noji, Christopher M. Yengo, Zhengyi Zhao and Ian Grainge. “Biological Nanomotors with a Revolution, Linear, or Rotation Motion Mechanism.” Microbiology and Molecular Biology Reviews 80 (2016): 161–186. Guo, Peixuan et al. “Controlling the revolving and rotating motion direction of asymmetric hexameric nanomotor by arginine finger and channel chirality.” ACS Nano 13 (2019): 6207–6223. Haque, Farzin & Geng, Jia & Montemagno, Carlo & Guo, Peixuan. (2013). Incorporation of viral DNA packaging motor channel in lipid bilayers for real-time, single-molecule sensing of chemicals and doublestranded DNA. Nature Protocols 8: 373–392. doi: 10.1038/nprot.2013.001. Haque, Farzin et al. “Methods for single-molecule sensing and detection using bacteriophage Phi29 DNA packaging motor.” Methods in Molecular Biology (Clifton, N.J.) 1805 (2018): 423–450. doi:10.1007/978-1-4939-8556-2_21. Ji, Zhouxiang, Michael Jordan, Lakmal Jayasinghe and Peixuan Guo. “Insertion of channel of phi29 DNA packaging motor into polymer membrane for high-throughput sensing.” Nanomedicine: Nanotechnology, Biology, and Medicine 25 (2020): 102170. Ji, Zhouxiang, Xinqi Kang, Shaoying Wang and Peixuan Guo. “Nano-channel of viral DNA packaging motor as single pore to differentiate peptides with single amino acid difference.” Biomaterials 182 (2018): 227–233. Jing, Peng, Farzin Haque, Anne P. Vonderheide, Carlo D. Montemagno and Peixuan Guo. “Robust properties of membrane-embedded connector channel of bacterial virus phi29 DNA packaging motor.” Molecular BioSystems 6(10) (2010): 1844–1852. Liang, Chenxi, Tao Weitao, Lixia Zhou and Peixuan Guo. “Translation of the long-term fundamental studies on viral DNA packaging motors into nanotechnology and nanomedicine.” Science China Life Sciences 63(8) (2020): 1103–1129. Schwartz, Chad et al. “The ATPase of the phi29 DNA packaging motor is a member of the hexameric AAA+ superfamily.” Virology 443(1) (2013): 20–27. doi:10.1016/j.virol.2013.04.004. Wang, Shaoying, Farzin Haque, Piotr Rychahou, B. Mark Evers and Peixuan Guo. “Engineered nanopore of Phi29 DNA-packaging motor for real-time detection of single colon cancer specific antibody in serum.” ACS Nano 7(11) (2013): 9814–9822. Wang, Shaoying, Zhengyi Zhao, Farzin Haque and Peixuan Guo. “Engineering of protein nanopores for sequencing, chemical or protein sensing and disease diagnosis.” Current Opinion in Biotechnology 51 (2018): 80–89. Wang, Shaoying, Zhi Zhou, Zhengyi Zhao, Hui Zhang, Farzin Haque and Peixuan Guo. “Channel of viral DNA packaging motor for real time kinetic analysis of peptide oxidation states.” Biomaterials 126 (2017): 10–17. Wang, Shaoying et al. “Three-step channel conformational changes common to DNA packaging motors of bacterial viruses T3, T4, SPP1, and Phi29.” Virology 500 (2017): 285–291. doi:10.1016/j.virol.2016.04.015.

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Part I Fundamental Mechanism of Biomotor Action

1

Biological Nanomotors with Linear, Rotation, or Revolution Motion Mechanism Peixuan Guo University of Kentucky

Hiroyuki Noji The University of Tokyo

Christopher M. Yengo Pennsylvania State University

Zhengyi Zhao Markey Cancer Center University of Kentucky

Ian Grainge University of Newcastle

CONTENTS 1.1 1.2

Introduction .............................................................................................................................. 4 Classification of Biomotors.......................................................................................................4 1.2.1 Rotation Motors ............................................................................................................ 5 1.2.2 Revolution Motors ........................................................................................................ 8 1.2.3 Linear Motors: Myosin, Kinesin, and Dynein ............................................................ 14 1.3 Structure of Biomotors ........................................................................................................... 15 1.3.1 Some Motor Components Display Hexameric Arrangements ................................... 15 1.3.2 Motor Structural Frame .............................................................................................. 16 1.3.3 Channel, Pore, or Surrounding Ring .......................................................................... 17 1.3.4 Factors for Distinction of Revolution Motor and Rotation Motors............................. 17 1.4 Motion Mechanism .................................................................................................................20 1.4.1 Energy Conversion: Transition among Entropy, Randomness, Affinity, and Conformation Change as Driving Force ...............................................20 1.4.2 Mechanism of Rotation Motors .................................................................................. 21 1.4.3 Mechanism of Revolution Motors ..............................................................................24 1.4.4 Mechanism of Linear Motors ..................................................................................... 29 1.4.5 Mechanism in Control Sequential and Coordination among Channel Subunits ........ 32 1.5 Potential Motor Applications .................................................................................................. 33 1.6 Concluding Remark and Perspectives .................................................................................... 33 Acknowledgments............................................................................................................................34 Competing Interests .........................................................................................................................34 References ........................................................................................................................................34 DOI: 10.1201/9780429203367-2

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4

1.1

Biomotors and their Nanobiotechnology Applications

INTRODUCTION

Electric motors impact almost all the aspects of daily life. Electric motors are electrical machines that convert electrical energy into mechanical energy, and further into kinetic energy to drive the operation of other devices. The first electric motor was created in the 1740s as a simple electrostatic device by the Scottish monk Andrew Gordon [1]. The principle of electric motors operations is based on Fleming’s left-hand and right-hand rules: when current flows through a wire that is placed in an external magnetic field, the wire will experience a force perpendicular both to the direction of the current flow and to the field. Similarly, bionanomotors, miniscule protein machines that produce mechanical motion by converting an energy source into work, are responsible for most forms of biological motion in living beings. These bionanomotors are essential in all aspects of crucial cellular processes critical to survival, such as mitosis, DNA replication, DNA repair, homologous recombination, Holliday junction resolution, RNA transcription, ATP synthesis, muscle contraction, viral genome packaging, and directional motility of cellular components to their destinations. Motor complexes allow the occurrence of such thermodynamically unfavorable processes. Biological nanomotors are ubiquitous and were once classified into two categories: linear and rotation motors, until the revolution motor without rotation was found as a third type of biomotor and is found to be widespread in bacteria, eukaryotic viruses, and dsDNA bacteriophages (for animation, see http://nanobio.uky.edu/movie.html). Viral DNA-packaging motor has been believed to be a rotation motor for decades [2]. The finding of the twisting of the motor channel structure, as revealed by crystal studies of biomotors, has reinforced this popular notion of a rotation mechanism for many years. However, extensive studies have revealed that none of the motor components rotate to a significant degree during genome packaging [3–7]. The finding of revolution motion has solved many mysteries regarding the motor structure and mechanisms. This review will focus on recent findings of various aspects of motor, including chirality, stoichiometry, channel size, entropy and conformational change, and energy usage in a wide variety of motor proteins: FOF1 ATPase, bacterial flagellar, helicases, viral dsDNA-packaging motors, bacterial chromosome translocases, myosin, kinesin, and dynein. dsDNA translocases are used to illustrate how these features relate to the motor mechanism in energy conversion, directional control, and sequential action. Nature has elegantly evolved a revolution mechanism devoid of coiling and tangling during lengthy dsDNA genome transportation.

1.2

CLASSIFICATION OF BIOMOTORS

Although different biomotors may possess different structures and have their distinct roles in cellular functions, they all need to undergo conformational changes that exert motions. Based on their motion mechanisms, the biomotors are categorized into three classes: linear, rotary, and revolution motors (Figure 1.1) [8–16]. Revolution motion is newly reported motion that has been mis-considered as rotation motion. Rotation is the circular movement of an object around its own axis, resembling the Earth that rotates on its axis in a complete cycle every 24 hours. Revolution is the turning of an object around the second object, resembling the action of the Earth that revolves around the Sun one circle per year (Figure 1.1). Many motors assembled into hexamers, and use ATP to trigger conformational changes towards different functions. Typical linear motors are myosin, kinesin, and dynein [17–19]; typical rotation motors are FOF1 ATP synthase, helicases, and bacterial flagella [20–22]; typical revolution motors are genomic dsDNA translocases for bacterial chromosome segregation and viral dsDNA packaging [12,23]. The representatives in this category include the dsDNA translocases FtsK of Escherichia coli and SpoIIIE of Bacillus; the dsDNA-packaging motor of bacteriophages phi29, T3, T4, T7, P22, lambda, SPP1; and the dsDNA-packaging motor of the large animal dsDNA viruses.

Biological Nanomotors: Linear, Rotation, or Revolution Mechanism

5

FIGURE 1.1 Illustration of different categories of motor. (a) Linear motors. (b) Rotation motors like the Earth that is rotating by its own axis. (Adapted from Ref. [49].) (c) Revolution motors resembling the Earth that is revolving around the Sun without self-rotation. (Adapted from Ref. [14] with permission from Elsevier.)

1.2.1 Rotation MotoRs FoF1 ATP synthase, helicases, and bacteria flagella are common representatives of this class of motors [20–22]. Besides the flagella, which is more intricate, many of these rotary ATPases are assembled into hexamers. Although many other nucleic acid-tracking ATPases are present in different oligomeric forms such as monomer and dimer, for those hexameric rotation motors that track alone nucleic acids, most, if not all, rotate along nucleic acids with one strand of DNA or RNA passing through the channel, while the other strand outside. Most rotation dsDNA translocases share the mechanisms that are distinct from the revolution dsDNA translocases. These rotation nanomotors include, but are not limited to, helicases [24–26], RNA polymerase [27], transcription termination factors [28], and others that participate in DNA recombination, repair, and Holliday junction resolution. RecA family ATPase monomers or dimers can assemble onto DNA as filament [29,30]; whether the functional unit in the complex is a hexamer helical open washer structure remains to be confirmed. FoF1 complex. FoF1 ATP synthase is found in the inner membranes of mitochondria, the thylakoid membranes of chloroplasts, and the plasma membranes of bacteria [31,32], responsible for ATP generation. This enzyme is composed of two rotary motors: Fo and F1. F1 is a membrane-protruding part and the catalytic domain of the ATP synthase. When isolated from the membrane, F1 acts as a rotary motor fueled with ATP hydrolysis. Fo is the membrane-spanning part of ATP synthase that conducts the proton translocation across the membrane down membrane potential. Upon the proton translocation, Fo rotates the ring-shaped rotor against the stator complex composed of the a and b subunits (Figure 1.2). Fo and F1 are connected by a common rotor axle and a peripheral stalk (Figures 1.2 and 1.3). These connections allow interconversion of proton motive force (pmf)1 and free energy of ATP hydrolysis via mechanical rotation of the subunit complex [20]. 1

The physicochemical term for pmf is electrochemical potential of proton.

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Biomotors and their Nanobiotechnology Applications

FIGURE 1.2 Structure of FoF1 ATP synthase and α3β3γ subcomplex of F1. (a) Reconstituted structure of FoF1 ATP synthase from crystal structures of isolated subunit or subcomplexes; α 3β3γε subcomplex (PDB code; 3OAA), δ (PDB code; 1ABV), b dimer (PDB code; 1B9U, 2KHK, 1L2P), c-ring (PDB code; 3UD0), and putative structure of the a subunit (PDB code; 1C17). Greenish parts represent the stator complex, including the peripheral stalk (δ-b2 subcomplex) that holds the α3β3 stator ring of F1 and ab2 stator of Fo. Brownish parts represent the rotor complex (γε-c-ring subcomplex). (b) Fo and F1 isolated from Figure 1.1a, both viewed from the top. (c) The original crystal structure of F1 from bovine mitochondria (PDB code; 1BMF). Sphere representations of the α, β, and γ subunits are shown in yellow, green, and red, respectively. Each β carries either AMP-PNP or ADP or none, and is designated as βATP, βADP, or βEmpty. (d) Conformational states of 3β subunits viewed from the side. α−β pairs are shown in green and yellow with the central γ subunit (red). α and β subunits are composed of the N-terminal domain, nucleotide-binding domain, and C-terminal domain (from bottom to top). βEmpty has an open conformation in which the α-helical C-terminal domain rotates upwards, opening the cleft of the nucleotide-binding pocket. Both βATP and βADP have a closed conformation entrapping the nucleotide within the closed pocket. All α subunits represent the open conformation.

Biological Nanomotors: Linear, Rotation, or Revolution Mechanism

7

FIGURE 1.3 Structure of Fo. (a) Crystal structure of c11-ring of Na+-transporting Fo from Ilyobacter tartaricus (PDB code; 1YCE). The blue spheres in the middle of the c11-ring represent bound Na+ ions. The stator ab2 complex is shown in the schematic drawing. The a subunit has two hemi-channels, each open to the periplasmic space or the cytoplasmic space. Proton transferring between the a and c subunits accompanies the rotation of the c-ring. Two c-subunit monomers at the interface of the a subunit are shown in red and green, respectively. (b) “Ion-locked” conformation of cGlu62 (yellow sphere representation) in the crystal structure of H+-transporting c15-ring from Spirulina platensis (PDB code; 2WIE). (c) “Ion-unlocked” conformation of cGlu59 (yellow sphere representation) in the crystal structure of H+-transporting c10-ring from yeast mitochondria (PDB code; 3U2F).

Bacterial flagella. Different species of bacterium use different motion methods in order to locate optimal growth conditions or avoid toxic substances. Most motile bacteria are driven by the rotation of flagellar filaments displaying a stiff helical structure with approximately 10 μm in length and 20 μm in diameter. Each filament has its own individual rotation motor at its base [21,22]. These motors can rotate clockwise to produce forward movement, or counterclockwise, resulting in tumbling and the changing of direction. The flagellar motor is powered by proton motive force. Many models have been proposed for the flagellar motor function, for example, the “electrostatic proton turbine” model [33–35] and the “turnstile” model [36,37]. Since many excellent updated reviews are available, readers are encourage to refer to Refs. [38–40]. RNA polymerase. The RNA polymerases are essential enzymes for the process of transcription. The RNA polymerase reads the DNA strand and transcribes it into RNA sequence. During this process, RNA polymerase rotates DNA through its channel, which has been shown by direct observation by the real-time optical microscopy measurement [41]. DNA helicase. Nucleic acid helicases comprise a class of enzymes that convert chemical energy into mechanical work for unidirectional translocation along nucleic acid and separate the nucleic acid duplexes into transient single-stranded intermediates. Helicases are involved in many cellular

8

Biomotors and their Nanobiotechnology Applications

processes, including DNA repair, replication, and recombination, and RNA transcription, remodeling, splicing, and translation. There are numerous ways to classify nucleic acid helicases. Based on the sequence, helicases are divided into six main groups (SF1–SF6). The cores of SF1 and SF2 enzymes are structurally similar to each other and contain similar sets of seven helicase signature motifs. The most well-studied SF3 helicase is the papilloma virus E1 helicase, and that of SF4 is T7 gp4 helicase (42). SF5 contains a bacterial Rho factor, which is an ATP-dependent hexameric RNA helicase involved in the termination of transcription in bacteria. SF6 contains some of the AAA+ proteins including MCM proteins as well as RuvB proteins [42].

1.2.2

Revolution MotoRs

DNA-packaging motor of dsDNA bacteriophages. By analogy to chromosome segregation or sporulation, packaging of a viral or bacteriophage genome into its capsid requires extensive translocation of the nucleic acid. It is perhaps not surprising that the motor proteins employed by many DNA viruses and phages belong to the FtsK-HerA family of translocases [43,44]. DNA-packaging motors of dsDNA bacteriophages have been historically classified as rotation motors, while extensive investigations have revealed that none of the motor components rotate during active motor actions (Figure 1.4) [3–5]. Connector rotation has been excluded both in phi29 motor using singlemolecule force spectroscopy in combination with single-molecule fluorescence polarization spectroscopy [4] and in T4 motor by tethering its connector for packaging assay [3] (Figure 1.4). In

FIGURE 1.4 Experiment of phi29 and T4 motors revealing that neither connector nor dsDNA rotation is required for active DNA packaging. (A) Direct observation of DNA packaging horizontally using a dsDNA with its end linked to a cluster of magnetic beads for stretching the DNA. “a” and “b” are the real-time sequential images of DNA-magnetic bead complexes. (Adapted from Ref. [5] with permission from the AIP publishing group.) (B) Experiment revealing that T4 motor connector does not rotate during packaging. The packaging activity is not inhibited with N-terminal of motor connector protein fused and tethered to its protease immune-binding site on capsid. (Adapted from Ref. [3] with permission from John Wiley and Sons.)

Biological Nanomotors: Linear, Rotation, or Revolution Mechanism

9

FIGURE 1.5 Schematic showing the sequential revolution motion in translocating dsDNA. (a) The binding of ATP to one ATPase subunit stimulates the ATPase gp16 to adopt a conformation with a higher affinity for dsDNA. ATP hydrolysis forces gp16 to assume a new conformation with a lower affinity for dsDNA, thus pushing dsDNA away from and transferring it to an adjacent subunit. Rotation of neither the hexameric ring nor the dsDNA is required. An animation is available at http://nanobio.uky.edu/movie.htm. (Adapted from Ref. [12]). (b) Diagram of Cryo-EM results showing offset of dsDNA in the channel of bacteriophage T7 DNA-packaging motor. The dsDNA did not appear in the center of the channel; instead, the dsDNA tilted towards the wall of the motor channel. (Adapted from Ref. [127] with the permission of the National Academy of Sciences.) (c) The revolution of dsDNA along the 12 subunits of the connector channel. (Adapted from Ref. [12].)

addition, tethering of DNA ends to bead clusters showed active DNA translocation with no observation of the rotation of the beads [5,7]. DNA was found to twist by as little as 1.5 degree per base pair translocated, confirming a non-rotation mechanism [6], since the 1.5°/bp  ×  10.5 bp/helical turn = 15.7° is far below the 360° per complete helical turn. This “rotation motors that do not rotate” puzzle was not solved until the breakthrough of the discovery of revolution motion in 2013 by Guo group [12–16] (Figure 1.5). Recent studies in bacteriophages suggested that the small twisting of the DNA is due to the conformational changes of DNA between A-form and B-form [45–47], and also the conformational changes of motor channel [48,49]. Such motors translocate DNA along the helix through unidirectionally revolution, resulting in a thermodynamic edge over rotation motors involving dsDNA translocation. The revolution motion is originally described in the well-studied phi29 dsDNA-packaging motor [12]. Viral dsDNA-packaging motors consist of a protein portal channel and two packaging components for packaging its genome into procapsid (Figure 1.6). The phi29 motor contains a dodecameric connector channel [50,51], a hexameric packaging RNA ring [7,52–54], and an ATPase gp16 hexameric ring [55,56] (Figure 1.6). All the dsDNA viruses known so far use a similar mechanism to translocate their genomic DNA into preformed protein shells, termed procapsids, during replication (see reviews [57–59]). More detailed descriptions and explanations about the motor structure characteristics and motion mechanisms can be found in the later sections.

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Biomotors and their Nanobiotechnology Applications

FIGURE 1.6 Depiction of the structure and function of phi29 DNA-packaging motor. (a) Side view of phi29 dsDNA-packaging motor (left) and top view of phi29 connector (right). (b) Hexameric pRNA generated from crystal structures of its 3WJ core and the AFM images of loop-extended hexameric pRNA. (c) DNA revolve inside the connector channel by contact with each connector subunit in a 30° transition step for each contact. (a–c. Adapted from Ref. [12].)

Inspired by the revolution motion found in phi29 dsDNA-packaging motor, studies in the other dsDNA motors have been carried out, and it was found that the revolution mechanism is a common feature shared by all the dsDNA-packaging motors, evidenced by the results from both crystal structural and biochemical studies. Crystal structure analysis of the motor channel of SPP1 [60], T7 [61], HK97 [62], P22 [63], and phi29 [51] revealed the existence of an anti-chiral arrangement between their channel subunits and the dsDNA helices in all these motors. Besides, the packaged genome in many viruses has been found to spool free from rotation tangles inside the capsid [64–67]. In phi29, a toroid of the dsDNA has been shown by Cryo-electron microscopy (Cryo-EM) around the portal region (Figure 1.7) [68–71], representing the accumulation of individual revolving DNA processed by Cryo-EM during revolution motion. Besides, a compression mechanism found in the T4 DNApackaging motor [46,47] also agrees with the revolution mechanism. Overall, the lack of rotation of the DNA during packaging allows tight, ordered packaging of the DNA with little or no knotting or tangling that could impair the DNA injection step during subsequent infection cycles. DsDNA translocases FtsK/SpOIII E superfamily for bacterial chromosome segregation. The FtsK/SpoIIIE family of proteins are hexameric dsDNA translocases found in many bacterial species, and were also proposed to use a revolution mechanism for DNA translocation (Figure 1.8). FtsK/SpoIIIE is part of the larger FtsK-HerA family that is present throughout bacteria and archaea [43], and also contains the motor proteins of various conjugative plasmids and transposons. Based on a core with a RecA-like fold, this family has some added features that set it apart from other RecA-like proteins [43]. FtsK is a dsDNA motor protein involved in the transportation of DNA and separation of intertwined chromosomes during cell division [72,73]. It functions to coordinate chromosome segregation, unlinking, and recombination with cell division so that the closing division septum is free from DNA [74], and it is one of the fastest and most powerful motors discovered to date, with a translocation speed of 17.5 kbps/s at 37°C [75] and a stalling force of 60 pN [76]. The FtsK motor consists of three functional components: one for DNA translocation, one for orientation control, and one for anchoring itself to the bacterial membrane (Figure 1.9) [77]. The C-terminal

Biological Nanomotors: Linear, Rotation, or Revolution Mechanism

11

FIGURE 1.7 Spooling of DNA within capsids of phages to support the revolution mechanism. The DNA spooling inside the capsids is shown using example of (a) phi29 bacteriophage. (Adapted from Refs. [68,69] with permission from Elsevier.) (b) P22 bacteriophage. (Adapted from Ref. [64] with permission from The American Association for the Advancement of Science.) (c) T7 bacteriophage. (Adapted from Ref. [70] with permission from The American Society for Biochemistry and Molecular Biology and Adapted from Ref. [71] with permission from The American Association for the Advancement of Science.) The toroid formed at the phi29 portal position might be an accumulation of the images of the revolution motion during packaging as shown in the image in the center. (Adapted from Ref. [15] with permission from Elsevier.)

DNA translocation motor component of FtsK can be further subdivided into three domains, namely, α, β, and γ [78]. While the α subdomain fold is unique to the FtsK family, the β subdomain, which contains residues for binding and hydrolyzing ATP, classifies FtsK both as a P-loop NTPase and as a hexameric translocase/helicase due to its RecA-like fold and sequence conservation common in these families [43]. The third subdomain, γ, has two distinct roles: it acts as a protein-protein interaction domain to activate Xer-mediated recombination at dif [79], and also acts as a DNA binding domain to recognize and bind to specific 8-bp DNA sequences on the chromosome. FtsK orienting polarized sequences (or KOPS), with the sequence GGGNAGGG [80, 81] are highly skewed in their distribution so that on each chromosome arm, the sequences are directed towards the terminus region, specifically switching orientation at the dif site. The role of this 8-bp sequence is to “determine” the directionality of translocation by acting as a recognition/preferential loading site for FtsK. Three γ domains bind to each 8-bp KOPS sequence (Figure 1.9c), and this leads to loading of an active hexameric to one side of the KOPS sequence (82) (Figure 1.10). This ensures not only that the motor is loaded correctly onto DNA in a specific orientation, but also that it is always translocated in one direction, which is towards the XerCD-dif site. Directional translocation of DNA is, therefore, sequence dependent; further, a translocating FtsK appears to ignore further KOPS sequences and reads through them [75,82]. SpoIIIE is an FtsK orthologue in B. subtilis that is vital for sporulation [83]. Like FtsK, SpoIIIE shares a conserved C-terminal domain that harbors three subdomains, namely, α, β, and γ [84,85]. The α and β subdomains translocate DNA through ATP binding and DNA-dependent hydrolysis [86], and the γ subdomain recognizes specific DNA sequences to guide DNA translocation [82,87]. These subdomains assemble into a hexameric ring that accommodates dsDNA in its central channel [78]. During spore formation, an asymmetric cell division occurs, and a complete chromosome copy is pumped into the smaller spore compartment from a larger “mother cell” using the ATPase-driven motor of SpoIIIE [83,88]. Recent reports proposed that SpoIIIE motors also use a revolution mechanism for DNA translocation [49]. Whether FtsK and SpoIIIE in fact respond to their respective

12

Biomotors and their Nanobiotechnology Applications

FIGURE 1.8 Illustration showing FtsK that may undergo the revolution mechanism. (a) One strand of the dsDNA contact with the inner channel wall of the hexameric ATPase. The continuous contact between DNA and ATPase does not require any rotation of the ATPase or DNA. (b) Each of DNA contact is expected to be separated by 60° along the inner surface of the ATPase hexameric channel. (c) Sequential action of dsDNA translocation. DNA is shown as a line. T represents for ATP-bound and D for ADP-bound (a–c. Adapted from Ref. [23] with permission from John Wiley and Sons.) (d) Speculation for the segregation and translocation of the mimivirus genome into the capsid via revolution mechanism similar to FtsK. (Adapted from Ref. [15] with permission from Elsevier.)

directionality sequences differently or whether the two related proteins behave in a similar fashion remains to be seen. DNA-packaging motor of large eukaryotic double-stranded DNA viruses. Comparative genomic studies of large eukaryotic dsDNA viruses, such as mimivirus, vaccinia virus, and pandoravirus, showed a remarkable similarity between their motor components and that of the FtsK-HerA superfamily [43,44,89], indicating that these viruses might also undergo revolution motion during genome packaging, allowing translocation of large genomes with minimal rotation of the DNA.

Biological Nanomotors: Linear, Rotation, or Revolution Mechanism

13

FIGURE 1.9 The FtsK motor protein. (a) Cartoon representation of the E. coli FtsK protein domain structure. The N-terminal domain is in red with each transmembrane domain represented by a black box. The numbers above the cartoon represent the amino acid numbers of FtsK. The C-terminus is subdivided into α, β, and γ domains. (b) Two views of the hexameric FtsK motor protein structure; on the left is a side on view emphasizing the α and β domains. Bound nucleotide (ADP) is shown in black in a space-filling model. On the right is a view down the symmetry axis, viewed from the β domain side. (c) Structure of three gamma domains bound to a KOPS DNA, seen along the DNA axis (left) and from side on (right).

FIGURE 1.10 A model of the FtsK motor loaded at a KOPS site. The N-termini of the γ domains are located on one side of the complex where they would connect to the motor domains of FtsK. This leads to loading of the motor to one side of the KOPS site so that the motor is pointing in a defined direction. This gives the motor its subsequent directional translocation (black arrow denotes the direction the motor would move along the DNA).

14

1.2.3

Biomotors and their Nanobiotechnology Applications

lineaR MotoRs: Myosin, Kinesin, and dynein

Myosin, kinesin, and dynein are linear-acting cytoskeletal motors. They are able to work in ensembles to generate large forces, such as during muscle contraction, or as single molecules to transport cargo along a specific cellular track. While myosins utilize actin filaments as tracks, kinesin and dynein are microtubule-based motors. The cytoskeletal motors are large protein families that are ubiquitously expressed in eukaryotic cells and are organized into classes based on their structure and function. Myosin motors are divided into 35 classes [17], while there are 14 classes of kinesin [90] and 9 classes of dynein [19]. The “linear motors” section of this review will focus on the myosin as an example of linear motion, while the reader is referred to excellent reviews for the kinesin [18] and dynein [19] mechanisms. Of the cytoskeletal linear motors, kinesin and myosin are more structurally related as their nucleotide-binding regions are similar to G-proteins and other P-loop ATPases. The key features of the active site are the conserved switch I, switch II, and P-loop (Walker A motif), which coordinate nucleotide binding and hydrolysis and also transmit key structural changes to the force-generating element of the motor [91] (Figure 1.11). The family of P-loop NTPases, G-proteins, kinesins, and myosins are thought to have evolved from a common ancestor [92]. Although the motor domains of the myosin and kinesin families are highly conserved, the N- and C-terminal domains are quite variable and allow dimerization, filament formation, protein-protein interactions, cargo binding, and light chain binding. Dyneins are members of the AAA+ family of ATPases, and their structure consists of a ring of six AAA+ ATPase modules [19]. The first AAA+ module is the key site of ATP hydrolysis, while sites 2–4 are also capable of hydrolyzing ATP, and sites 5 and 6 are not capable of hydrolyzing ATP. Dynein also contains a stalk and microtubule-binding domain as well as a cargo-binding tail that extends from the AAA+ ring.

FIGURE 1.11 A simplified schematic representation of the ATPase cycle of myosin. The proposed mechanism of how the motor is primed (recovery stroke) and generates force by coupling movement of the lever to key steps in the ATPase cycle.

Biological Nanomotors: Linear, Rotation, or Revolution Mechanism

1.3 1.3.1

15

STRUCTURE OF BIOMOTORS soMe MotoR CoMponents display HexaMeRiC aRRangeMents

Many members of the ASCE superfamily function with hexameric assemblies [93–96]. However, structural studies of viruses once led to the popular fivefold/sixfold mismatch mechanism [2] in 1978 based on the findings that viral icosahedron capsid is composed of many pentamers and hexamers [97] and that the DNA-packaging motor of dsDNA viruses resides within the pentameric vertex [98] with a dodecameric motor channel [61,99–103]. In 1998, the pRNA ring was first determined to be a hexameric ring [53,104] (featured by Cell [105]). Some future studies by others have shown a pentameric model [106–108]; the hexameric pRNA formation was verified by Cryo-EM [109], biochemical analysis [53,104,105], single-molecule photobleaching study [7], gold labeling imaging by electron microscopy (EM) [110,111], and RNA crystal structure studies [54]. The observation that all six copies of pRNA remain at the active motor during packaging [7] (Figure 1.12) excluded the assumption that the motor components change from hexamer to pentamer when the packaging initiates [108,112,113]. More recently, the ATPase gp16 of phi29 dsDNA-packaging motor, which belongs to the ASCE superfamily, has also been confirmed to be a hexamer as its

FIGURE 1.12 Single-molecule photobleaching assay confirms the presence of six copies of phi29 motor pRNA. (a) pRNA dimer design constructed with Cy3- and Cy5-pRNA. (b) Comparison of empirical photobleaching steps with theoretical prediction of Cy3-pRNA in procapsids bound with dual-labeled dimers. (c) Photobleaching steps of procapsids reconstituted with the dimer. (a–c. Adapted from Ref. [7] with permission from John Wiley and Sons.)

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Biomotors and their Nanobiotechnology Applications

FIGURE 1.13 Stoichiometric assays showing the formation of the phi29 ATPase hexamer. (a) Native gel revealed six oligomeric states of the ATPase; the hexamer formation increases as the concentration of protein increases. (b) Slab gel showing the binding of ATPase to dsDNA in a 6:1 ratio, imaged in GFP (upper) and Cy3 channels (lower) for ATPase and dsDNA, respectively. (c) Quantification by varying the molar ratio of [ATPase]:[DNA]. The concentration of bound DNA plateaus at a molar ratio of 6:1. (a–c. Adapted from Ref. [55].)

final oligomeric state (Figure 1.13) [55], as demonstrated by binomial distribution analysis, qualitative DNA binding assays, capillary electrophoresis (CE) assays, and electrophoretic mobility shift assays (EMSA) [55,114,115]. Besides, the four steps of motor motion reported by the biophysical studies (Figure 1.14a) have been shown to be related to the four relaying lysine layers embedded inside the phi29 connector channel inner wall (Figure 1.14b and c) [13], further evidence against the pentamer model [107,116]. Assembly of a hexamer is also crucial to the motor function, for example, the ATPase function of the FtsK (and SpoIIIE) DNA motors. The α and β domains hexamerize to form DNA translocase motor, a ring-shaped multimer with a central channel through which the dsDNA substrate is threaded [78]. EM of FtsK(C) has revealed a DNA-dependent hexamer formation and DNA passage through the hexameric ring. α and β each form a hexameric ring, and the two rings sit atop one another with a ~10Å cleft separating them and two strands connecting the rings (Figure 1.9b).

1.3.2

MotoR stRuCtuRal FRaMe

Molecular motors exhibit a wide variability in structure. Most are quite complex, but in general, the structures consist of the same essential components: a mechanical frame that consists of both moving and static parts, along with an energy supply. The mechanical frame is formed by proteins in most molecular motors, though the DNA-packaging motor of bacteriophage phi29 also contains

Biological Nanomotors: Linear, Rotation, or Revolution Mechanism

17

FIGURE 1.14 Four steps of pauses for each circle during the packaging of phi29 dsDNA. (a) The presence of four lysine residues of motor channel protein leads to the formation of four positively charged rings in different motors. (Adapted from Refs. [49,163] with permission from Elsevier.) (b) Diagram showing DNA revolution inside phi29 connector channel with four steps of pause due to the interaction of four positively charged lysine rings with the negatively charged dsDNA phosphate backbone. DNA revolution across the 12 connector channel subunits is shown. (Adapted from Ref. [15] with permission from Elsevier). (PDB codes: Phi29-gp10, 1H5W; SPP1-gp6, 2JES; P22-gp1, 3LJ5.)

a hexamer of RNA molecules as an essential component [52]. The DNA RNA polymerases are track-laying motors. DNA and RNA can be considered components of these motors since they are required for the unwinding or polymerization process [117–120]. The structure of the motor domain, which contains all the elements capable of converting chemical energy into mechanical work, is highly conserved within the various classes of motors [121,122].

1.3.3

CHannel, poRe, oR suRRounding Ring

During cell segregation or binary fission, circular closed dsDNA can be translocated by the dsDNA translocases without breaking the covalently bonded backbones [77,78,88]. Besides, concatemer dsDNA is the packaging substrate in many dsDNA viruses [123–125]. This process can be finished by the dsDNA translocation motor, since the ATPase monomers can assemble into a hexamer on the DNA itself without a free 5′ or 3′ dsDNA end utilizing the energy from ATP binding [49]. This is confirmed by the single-molecule fluorescent imaging and EM imaging, showing that the first step in phi29 DNA packaging is the binding of multiple gp16 in a queue along the dsDNA, evidenced by the observation of a string of multiple Cy3-gp16 complexes on dsDNA chains in the presence of non-hydrolyzable ATPγS [12].

1.3.4

FaCtoRs FoR distinCtion oF Revolution MotoR and Rotation MotoRs

Among these three types of motors, the differentiation of rotation motor from revolution motor is relatively challenging, especially with the fact that dsDNA translocation motors involve both rotation and revolution motors. Below, we will introduce two simple ways to distinguish these two types of motor. Chirality: left-handed for revolution motor and right-handed for rotation motor. Chirality is one of the criteria to distinguish revolution motors from rotation motors. The motor channels

18

Biomotors and their Nanobiotechnology Applications

FIGURE 1.15 The use of channel chirality to distinguish revolution motors from rotation motors. (a) In revolution motors, the right-handed DNA revolves within a left-handed channel. (b) In rotation motors, the right-handed DNA rotates through a right-handed channel via the parallel thread, with RecA and DnaB shown as examples. (Adapted from Ref. [49].) (PDB codes: RecA, 1XMS; P22-gp1, 3LJ5).

(connector) of SPP1 [60], T7 [61], HK97, P22 [63], and phi29 [51] all adopt an anti-chiral arrangement between the left-handed motor connector subunits and the right-handed DNA helices during packaging (Figure 1.15) [12,13]. No apparent homology has been shown among these portal proteins by sequence alignment. However, they all share very similar three-dimensional structures with conserved regions, and a highly similar pattern with particularly a sequence of α-β-α-β-β-α stretches. The left-handed helices of each of the 12 subunits lining inside the channel all tilt at a 30° angle (Figure 1.16), locating at the same conserved sequence at the last alpha helix of the α-β-α-β-β-α stretches, indicating the critical role of this conserved 30° anti-chiral arrangement in dsDNA packaging [49]. Such anti-chiral structure greatly facilitates the controlled motion of one-way packaging of dsDNA during its revolution action with no rotation, coiling, or torsion force by the contact between each of the 12 connector subunits with a 30° transition for each contact [12]. On the other hand, rotation motors use the right-handed channels for the right-handed dsDNA translocation as parallel thread. Channel size: larger than 3 nm for revolution motor and smaller than 2 nm for rotation motor. Rotation motors mostly process one of the nucleic acid strands that is threaded through the center of the channel, and the other is excluded during translocation [126]. Rotation motors adopt a channel with full contact with nucleic acid during its translocation through the center of the channel; as a result, the channel size has to be close to or smaller than the diameter of one strand of dsDNA, which is smaller than 2 nm. However, for revolution motors, both strands of the dsDNA are advancing within the channel through its contact with the channel wall from the side, instead of proceeding through the center of the channel, in agreement with the Cryo-EM images showing that the T7 dsDNA core tilts relatively to its channel axis. A counterclockwise motion of the dsDNA in T7 motor was observed viewing from the N-terminus of T7 connector [127] (Figure 1.5b), in agreement with the direction of dsDNA revolution in phi29 dsDNA-packaging motor [12,13,59,128] (Figure 1.5a). The revolution motor channels are thus much larger than the dsDNA in order for the sufficient room for revolution. Crystal structures of different motors have confirmed the channel sizes, confirming that the diameters of the revolution motor channels are larger than 3 nm, while those of the rotation motors are smaller than 2 nm [49], as evidenced in revolution motors of phi29, SPP1, T4, T7, HK97, and FtsK, and rotation motors of Rho factor, E1 helicase, TrwB, ssoMCM, and RepA [49] (Figure 1.17).

Biological Nanomotors: Linear, Rotation, or Revolution Mechanism

19

FIGURE 1.16 Quaternary structures showing the presence of the left-handed 30° tilting of the connector channel of different bacteriophages. (a) The external view and (b) the cross section view of the motor, showing the anti-parallel configuration between the left-handed connector subunits and the right-handed dsDNA helices. The 30° tilt of the helix (highlighted) relative to the vertical axis of the channel can be seen in a cross-sectional internal view of the connector channel and the view of its single subunit as shown in b. (a–b. Adapted from Refs. [15,49] with permission from Elsevier.) (PDB IDs: Phi29-gp10, 1H5W; HK97 familyportal protein, 3KDR; SPP1-gp6, 2JES; P22-gp1, 3LJ5; T7-gp8 EM ID: EMD-1231).

20

Biomotors and their Nanobiotechnology Applications

FIGURE 1.17 Comparison of the size of channels between biomotors using rotation mechanism (a) and biomotors using revolution mechanism (b). The revolution motors use larger channels (> 3 nm in diameter), while rotation motors use smaller channels (< 2 nm in diameter) during DNA translocation. (Adapted from Ref. [49].) (PDB codes: RepA, 1G8Y; TrwB: 1E9R.; ssoMCM, 2VL6; Rho, 3ICE.; E1, 2GXA; T7-gp4D, 1E0J; FtsK, 2IUU.; Phi29-gp10, 1H5W; HK97 family-portal protein, 3KDR; SPP1-gp6, 2JES; P22-gp1, 3LJ5; T7-gp8 EM ID: EMD-1231).

1.4 MOTION MECHANISM 1.4.1

eneRgy ConveRsion: tRansition aMong entRopy, RandoMness, aFFinity, and ConFoRMation CHange as dRiving FoRCe

It has been well established that biomotor motion can be coupled to both ATP binding and ATP hydrolysis [129,130]. Several assays were used to study the ATP-ATPase interaction and elucidate how ATP energy translates to physical motion in the phi29 DNA packaging. EMSA suggests that the ATPase undergoes conformational entropy changes upon ATP binding or ATP hydrolysis, leading to a high or low binding affinity of ATPase towards DNA, respectively [12,55,131]. These changes in ATPase force the substrate DNA away from the ATPase and lead to physical motion of genomic DNA towards the second subunit, thereby moving the DNA towards the interior of the viral protein shell (Figure 1.18). Six ATP are used to complete one revolution cycle, with one ATP to package 1.75 bp of dsDNA (10.5bp/6ATP = 1.75 bp/ATP). Such conformational changes are abolished by the site-directed mutation to the Walker A motif [55], which has been identified [56] and confirmed [12,56,132] to be responsible for ATP binding. The mutation to gp16 Walker B motif that is required for ATP hydrolysis eliminates the catalytic force step [55]. Thus, the conformational entropy change of the ATPase is a process that couples motion with the free energy changes, and most likely, an intrinsic property of the protein through evolution. The P-loop lysine, glutamic acid of general base, and arginine finger are well known to be critical to retain the catalytic power of ATPases. P-loop lysine is the lysine in the highly conserved phosphate-binding loop (P-loop) among NTPases, of which common sequence is GXXXXGKT/S. The glutamic acid, thought to be a general base, is also well conserved among ATPases that interacts the γ phosphate via a water molecule. The glutamic acid general base has been thought to

Biological Nanomotors: Linear, Rotation, or Revolution Mechanism

21

FIGURE 1.18 Model of sequential mechanism of sequence action of phi29 DNA-packaging motor. Binding of ATP to the conformationally disordered ATPase subunit stimulates an entropic and conformational change of the ATPase, thus fastening the ATPase at a less random configuration. This lower entropy conformation enables the ATPase subunit to bind dsDNA and prime ATP hydrolysis. ATP hydrolysis triggers the second entropic and conformational change, which renders the ATPase into a low affinity for dsDNA, thus pushing the DNA to the next subunit that has already bound ATP. These sequential actions promote the movement and revolution of the dsDNA around the hexameric ATPase ring. (Adapted from Ref. [15,49] with permission from Elsevier.)

activate the intervening water molecule to induce nucleophilic attack of the water molecule on the γ phosphate. Recent quantum chemical calculation and single-molecule analysis revealed that the role of general base is not water activation but the enhancement of proton transfer from phosphoester to bulk solution [133]. In some ATPases, the glutamic acid also plays a role as a sensor for ATP-binding state, and changes orientation drastically upon ATP binding [134]. The arginine finger is also highly conserved among G proteins, AAA proteins, and RecA-type ATPases. The arginine finger is located at an interface of nucleotide-binding subunits; ATP binds the interface of two subunits, one of which possesses most of ATP-binding residues, while the other provides the arginine finger. The crystal structure of bovine mitochondrial F1 (MF1) with chemical analog of the γ phosphate suggests the arginine finger stabilizes transient state of hydrolysis reaction that was supported by biochemical [135], theoretical [136], and single-molecule studies [137]. The AAA+ family also contains another arginine, in a structurally separate location to the arginine finger, that acts to couple ATP binding and hydrolysis to conformational changes between subunits; this has been termed the Sensor II motif [138,139].

1.4.2

MeCHanisM oF Rotation MotoRs

Step rotation of F1. Rotation of the isolated F1 motor driven by ATP hydrolysis was directly observed with an optical microscope [32,140]. Since the unidirectional rotation was first visualized for F1 from Bacillus PS3, several kinds of rotary ATPases, F1’s from Escherichia coli [141], chloroplast from spinach [141], human mitochondria (T. Suzuki, personal communication), and V-ATPases from Thermus thermophiles [142] and Enterococcus hirae [143]), were examined in the rotation assay. Among them, F1 from Bacillus PS3, termed TF1 (thermophilic F1), is the best characterized

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Biomotors and their Nanobiotechnology Applications

in terms of rotary dynamics. Therefore, the rotation features of F1 introduced hereafter are based on the findings in the single-molecule rotation assay of TF1 unless mentioned. In consistent with the pseudo-3-fold symmetry of the α3β3 stator ring, F1 rotates γ in discrete 120° steps [144], each coupled with a single turnover of ATP hydrolysis [144,145]. An intermediate state was also observed after ATP binding and before hydrolysis at an 80° angle from the ATP-waiting angle [146]. Thus, each 120° step can be resolved into 80° and 40° substeps (Figure 1.19). A recent reaction scheme suggests that 80° substep is driven by ATP binding and ADP release that occur on different β subunits and that the 40° substep is initiated after the hydrolysis of bound ATP and Pi release that also occur on different β subunits [147] (Figure 1.19c). Single-molecule rotation assay of F1. Dynamic behavior of ATP-driven rotation of F1 is well characterized in the single-molecule rotation assay where the α3β3 stator ring is immobilized on a coverslip and the rotation probe is attached onto the outwardly protruding part of the γ subunit of F1 (Figure 1.19a). In earlier studies, a fluorescently labeled actin filament (0.5–5.0 m) was used [140,144]. In recent studies, nanoparticles with high scattering coefficient are often used, such as polystyrene bead [148], gold nanocolloid [148], and nanorod [149,150], which enable imaging with high spatiotemporal resolution. In order to manipulate the rotation of F1, submicron-sized magnetic beads are attached on the γ subunit as a rotation probe as well as handle to control the orientation of the γ subunit with magnetic tweezers [145,151,152]. Conformational transition of the β subunit. Many lines of experimental data showed that the catalytic β subunit undergoes a large conformational change [153–155]. The clearest data came from the crystal structure of MF1 that showed the β subunit changes the conformational state upon nucleotide binding, rotating the C-terminal helical domain inwardly [153]. This conformation is termed closed form. On the other hand, nucleotide-free β subunit assumes an opened conformational state. It seems that when nucleotide-free β subunit undergoes opened-to-closed conformational transition upon nucleotide binding, it apparently pushes the γ subunit. Therefore, ATP binding is long thought to be a major torque-generating step among elementary catalytic steps: binding, hydrolysis, and release of ADP and Pi. nuclear magnetic resonance (NMR) study (154) and single fluorophore imaging [155], and recent-high speed atomic force microscopy (AFM) showed that the β subunit actually undergoes opened-to-closed conformational transition upon ATP binding [156] as expected from the crystal structure of MF1. Taking into account that the induced-fit process is a major torque-generating step, it was proposed that the progressive hydrogen bond formation between the phosphate moiety of bound ATP and the catalytic residues is the main driving force of the induced-fit-type conformational change: the opened-to-closed transition of the β subunit. The accompanying swing motion of the C-terminal domain of the β subunit would push the γ subunit to induce the rotation (Figure 1.20). PMF-driven rotation of Fo. The most straightforward approach to address the dynamic feature of c-ring rotation is imaging c-ring rotation under pmf-driven conditions. A pioneering work was done by Börsch’s group [157]. They employed single-molecule Förster resonance energy transfer (FRET) measurement to visualize c-ring rotation [158]. Donor and acceptor fluorescent dyes were introduced to the a subunit and one of the c subunit the stator and rotor subunits of EFoF1 reconstituted into liposomes. Under ATP synthesis condition, FRET signal showed multiple states that were attributed to intervening pause of c-ring at every 36° steps based on the tenfold symmetry of the c-ring from Escherichia coli. Following Börsch’s group, single-molecule rotation assay of FoF1 in ATP synthesis condition was reported [159], where EFoF1 molecules were reconstituted into a supported membrane and pmf was charged by uncaging the caged proton between coverslip and supported membrane. In this experiment, EFoF1 molecules showed clear 120° steps, suggesting that kinetic bottleneck was a catalytic event on F1. These experiments still have a critical experimental drawback that is the electrochemical potential which is transient and not stable that prevents the elaborated analysis. Thus, the single-molecule analysis on rotary dynamics of Fo still requires technical advancement for the elucidation of rotary dynamics. One remarkable technical achievement is the arrayed lipid bilayer chamber (ALBiC) system that displays a million of femtoliter chambers,

Biological Nanomotors: Linear, Rotation, or Revolution Mechanism

23

FIGURE 1.19 Single-molecule rotation of F1. (a) A schematic image of the experimental setup. The α3β3 ring is fixed on the glass surface. A probe (fluorescently labeled actin filament or 40-nm colloidal gold) is attached to the γ subunit. (b) (Left) Rotation of F1 with three binding pauses separated 120°, which is caused by slow ATP binding at 200 nM. Inset is the trajectory of the rotation. (Center) Rotation of a mutant F1 (βE190D) with three catalytic pauses at 2 mM ATP. Each pause is caused by the extremely slow ATP hydrolysis by the mutant. (Right) Rotation of mutant F1 (βE190D) at 2 µM ATP. Due to slow ATP binding and hydrolysis, 6 pauses are observed. The pauses before the 80° (arrowheads) and 40° (arrows) substeps correspond to binding and catalytic pauses, respectively. (c) Chemomechanical coupling scheme. Each circle indicates the chemical state of the catalytic sites. One catalytic site is highlighted in dark green. The central arrow (red) represents the angular position of the γ subunit. Each catalytic site retains the bound nucleotide as ATP until the γ subunit rotates 200° from the binding angle (0°). After a 200° rotation, the catalytic site executes the hydrolysis of ATP into ADP and Pi, each of which is released at 240° and 320°, respectively.

each sealed with a lipid bilayer patch [160]. This allows single-molecule measurement of transporting activity of membrane proteins such as alpha hemolysin and ATP synthase. One advantage of the ALBiC system is very hermetic sealing of chambers with lipid bilayers. Such new membrane technology would pave a way to access the single-molecule study of rotary ATPase as well as other membrane protein machines, by combining with single-molecule imaging techniques.

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Biomotors and their Nanobiotechnology Applications

FIGURE 1.20 The opened-to-closed transition of the β subunit. The accompanying swing motion of the C-terminal domain of the β subunit would push the γ subunit to induce the rotation.

Rotation of helicases. Several mechanisms of helicase translocation along nucleic acid lattices have been proposed. Currently, the most popular models are “inch-worm stepping mechanism” and “Brownian motor mechanism” [161,162]. Translocation begins with one binding site tightly bound to the substrate, while the other one bound weakly to it. “Inching forward” of the helicase is coupled with the NTP binding and hydrolysis cycle. Binding and coordination of the NTP-metal ion complex results in the large conformational change, which closes the cleft between the two nucleic acid-binding sites. NTP hydrolysis and subsequent release of the NDP and inorganic phosphate is followed by the reversal of the conformational change, which results in the relative movement of the domain (or subunit) containing leading nucleic acid-binding sites. One cycle of action is completed in six conformational changes. Brownian mechanism requires only one nucleic acid-binding site on the helicase, while two distinct conformational states were proposed with high or low affinities towards its substrate depending on the different NTP ligation states. Nucleic acids are translocated by the combination of Brownian motion and power stroke.

1.4.3

MeCHanisM oF Revolution MotoRs

One-way traffic of dsDNA-packaging motor. The special structure of the motor generates several factors that coordinate to drive the dsDNA with a one-way traffic mechanism. As mentioned in the previous sections, ATP binding induces an entropic and conformational force of the ATPase for high affinity to approach dsDNA. ATP hydrolysis induces a second entropic and conformational change, resulting in a low affinity for dsDNA and subsequently pushing dsDNA move forward towards the next subunit of ATP hexamer. Second, the 30° left-handed twist of the channel wall produced an anti-parallel arrangement between the channel wall and the right-handed helix of the dsDNA, resulting in a vector force. Third, the one-orientation flow loops within the channel found in SPP1, phi29, and T4 generate a vector force for the one-way trafficking of dsDNA (Figure 1.21).

Biological Nanomotors: Linear, Rotation, or Revolution Mechanism

25

FIGURE 1.21 Role of the flexible inner channel loop in DNA one-way traffic. (a) Flexible loops within the phi29 (left) and SPP1 (right) connector channels function to interact with DNA, facilitating DNA to move forward but blocking the reversal of DNA during DNA packaging. (b) Demonstration of one-way traffic of dsDNA through wild-type connectors using a ramping potential and by switching polarity (right). (c) SsDNA is translocated via two-way traffic with a loop-deleted connector. (Adapted from Ref. [13] with permission from American Chemical Society.)

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Biomotors and their Nanobiotechnology Applications

A mutant phi29 connector with the deletion of the internal loop N229-N246 failed to produce any virion [48,163–165]. Single-channel conductance assay also showed the one-way property for wildtype connector [13,128], while the two-way property for the loop-deleted connector (Figure 1.21) [13,166]. These results suggest that the channel loops act as a ratchet during DNA translocation to prevent the DNA from leaving, in line with the “push-through one-way valve” model [13,59,128,163]. Analysis of the crystal structure and Cryo-EM density map of SPP1 channel loops reveals that these loops are located in a close proximity to dsDNA via non-ionic interactions. The channel loops of phi29 and SPP1 might play similar roles in directional traffic of dsDNA to enter the capsid through the connector channel [167]. Besides, the 5ʹ to 3ʹ mode of revolution of one DNA strand along the channel wall to generate the vector force agrees with published data from T4 and phi29. During motor packaging, dsDNA is processed by contacting the connector with one strand of DNA in the 5ʹ to 3ʹ direction [168]; modification of phi29 genome DNA in the 5ʹ to 3ʹ direction strand was found to stop dsDNA packaging [169] (Figure 1.22). Extensions up to 12 bases at the end of the DNA can be tolerated; however, extensions to 20 or more bases significantly blocked the DNA packaging of the T4 motor [170]. Furthermore, the electropositive lysine layers present in many phages channels interact with one of the strand of the electronegative dsDNA phosphate backbone, resulting in a relaying contact that

FIGURE 1.22 Effect of DNA chemistry and structure on its packaging. (a) Demonstration of blockage of dsDNA packaging by single-stranded gaps. When a single-stranded gap is present, only the left-end fragment of phi29 genomic DNA is packaged. (Adapted from Ref. [168] with permission from Elsevier.) (b) Chemical modification of the negatively charged phosphate backbone on DNA packaging. Modification on the 3′→5′ strand does not block dsDNA packaging, but alternation on the other direction seriously affects DNA packaging. The results support the finding of the revolution mechanism showing that only one strand of the dsDNA interacts with the motor channel during revolution. (Adapted from Ref. [169] with permission from Nature Publishing Group.)

Biological Nanomotors: Linear, Rotation, or Revolution Mechanism

27

facilitates one-way motion and the generation of steps of transitional pausing during dsDNA translocation [12,13,106]. The negatively charged phi29 connector interior surface is decorated with 48 positively charged lysine residues, resulting in the formation of four 12-lysine rings, as revealed by the connector crystal analysis [51] (Figure 1.14). The lysine layers are nonessential for DNA packaging, while the interactions between the lysine and DNA backbone are involved in motor action. Similar patterns of four relaying electropositive lysine layers have been found in phage of SPP1, P22, and phi29 [51,113] within the predominantly negatively charged connector channel surface, which results in the uneven speed alternations during the DNA translocation process with four pauses [12,13,106] as previously reported in both phi29 [106,116] and T4 [171]. The effects of the lysine layers of the genome translocation can be interpreted from the structural aspects. Taking phi29 dsDNA-packaging motor as an example, its four lysine layers fall vertically within a 3.7-nm range [51] inside the 7-nm connector channel, spaced averagely ~0.9 nm apart. For B-type dsDNA, ~2.6 bp will advance through each rise between two lysine layers (0.9 nm/0.34 nm·bp−1 = ~2.6 bp). For every cycle of the DNA revolution (360°, 10.5 bp) through the channel of 12 subunits, a 0.875 mismatch shows up between the negative DNA phosphate base and the channel subunits with positive lysine layer (10.5/12 = 0.875). To compensate the distance variation due to this mismatch, the dsDNA phosphate backbone will interact with the positively charged lysine in the next subunit, leading to a slight pause in DNA advancement (Figure 1.14). The continuation of the interactions between lysine layers and DNA backbones leads to the four pauses mentioned above during packaging. dsDNA translocases FtsK/SpOIII E superfamily. The crystal structure of the hexameric FtsK motor domain from Pseudomonas aeruginosa revealed a sixfold symmetric ring with ADP bound in the active site of every subunit [78]. There was very little clue from this structure alone as to how the protein could convert the chemical energy from ATP hydrolysis into movement of the DNA substrate. However, the same study also presented a different crystal form of the motor domain, a monomeric form with ATP bound. A comparison of the two structures showed that when the β domains were superimposed, the α domains were shifted relative to each other in a hinge-like opening of α from β. The point in the α domain juxtaposed to the DNA substrate was able to move by 5.5 Å by this hinge-like mechanism (equivalent to 1.6 bp). If this movement were correlated with ATP hydrolysis, it would provide a mechanism by which the protein could pump DNA. The translocation of dsDNA with 1.6-bp subunit of FtsK [23,78] agrees well with the quantification in phi29 DNA-packaging motor that each ATPase subunit uses one ATP to package 1.75 nucleotides [12,13,16,55,56], and the bacteriophage T3 system with 1.8 bp per ATP [172]. Based on these data, an inchworm mechanism has been proposed for FtsK [23,78] with ATPase subunits acting in a sequential manner. In the model proposed by Massey et al., each ATP hydrolyzed moves the DNA by 2 bp (~6.8 Å, slightly larger than the 5.5 Å difference between the two crystal forms) in an inchworm-like movement with contacts between both domains with the central DNA. The relative strength of interaction between each domain and the DNA is dependent upon the ATP binding or hydrolysis state; upon hydrolysis of ATP, one DNA contact is lost, while the other contact is strong but is shifted by ~6.8 Å, resulting in the net movement of DNA along the central channel of 2 bp. An integer number of bases was chosen as it allows each monomer in the ring to contact the dsDNA in the same manner at every subunit, whether this be contact with the repeating sugar-phosphate backbone, or with the bases themselves. Movement by a non-integer number of bases would mean that the protein-DNA contact necessary for the movement of the substrate would be different in any two adjacent subunits. The movement of dsDNA by one subunit also functions to bring the next monomer in the ring into register with the DNA, and so hands the DNA substrate on to the adjacent monomer with minimal rotation (Figure 1.8b). The next subunit then also translocates 2 bp of DNA upon hydrolyzing ATP, and so on around the ring. The result is 12 bp of DNA translocated per catalytic cycle of hexamer where all six monomers hydrolyze ATP once. This figure is close to the 10.5 bp per helical turn in B-form DNA, with the extra 1.5 bp being manifested as a slight twisting. If the protein is anchored (both FtsK and SpoIIIE and membrane bound at their N-termini), then this results in the generation of DNA supercoiling: +ve supercoiling ahead of the motor and –ve

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Biomotors and their Nanobiotechnology Applications

supercoiling behind. Indeed, the induction of small supercoiling has been seen in bulk biochemical assays [173] and in single-molecule experiments [174]. The observed supercoiling induction for FtsK translocation of one +ve supercoil ahead of the motor per 150 bp translocated is in broad agreement with the rotary inchworm model proposed. The inchworm model also proposed an obligatory hand-off event between adjacent monomers within a single ring, such that the presence of a single catalytically inactive subunit would effectively inactivate the entire hexamer [78]. This was backed up by biochemical data: mutants which were unable to bind ATP were mixed in different ratios with wild-type subunits and the relative ATPase activity was compared to hexamers consisting of only wild-type subunits. With increasing amount of mutant subunits, the ATPase activity decreased rapidly, closely following the predicted pattern for positioning of the hexamer by an inactive subunit. This agrees with the finding that the incorporation of one inactive subunit of the phage phi29 DNA-packaging motor completely blocks the function of the ATPase ring [12,13,15,55]. Interestingly, a fusion protein has been produced in which three motor domains were joined to each other in a single polypeptide, effectively a covalent trimer of FtsK [175]. This construct was found to be a very active DNA translocase motor. Within this trimeric construct, the Walker A and Walker B motifs, for nucleotide binding and hydrolysis, respectively, could be mutated, and this led to the surprising finding that a single active-site mutant, or two non-adjacent mutants per hexamer, did not cause a great decrease in ATPase activity and did not significantly decrease the speed of translocation along dsDNA [175]. However, the presence of these mutations did reduce the ability of the hexamer to produce force as judged by the ability of the protein to displace either protein or DNA triplexes. It is important to consider that when the ATPase subunits were fused into concatemer, there may be unknown and unintended alterations either to the ATPase activity of individual monomers or to their ability to form higher-order multimers that could mask the effect of mutant subunits. Conversely, a number of single-molecule studies have now shown that the linked trimeric FtsK proteins appear to translocate and respond to nucleotide similarly to unlinked hexamers [75,176]. Nevertheless, in order to explain these results, a new model was proposed in which more than one subunit within a hexameric ring would contact DNA concurrently. This model was based upon the escort or “spiral staircase” model of Rho and E1 helicases [26,177]. In these hexameric helicases, multiple subunits can contact the DNA/RNA substrate at the same time, with the single contact point for each monomer being at a different level around the ring, rather like the stairs in a spiral staircase. ATP hydrolysis moves one of the contact points downwards through the ring forcefully, and the other contacts move along passively. When the last contact point at the bottom of the ring is reached, the protein arm becomes free and can then move back up the top to re-engage with the polynucleotide substrate and begin the cycle of movement down the staircase again. With a flexible and compressible single-stranded DNA/RNA substrate, the movement of the protein-DNA contacts is small enough that the protein can maintain with the DNA/RNA through a full catalytic cycle of the every subunit in the ring. However, dsDNA is a much stiffer and non-compressible substrate than ssDNA or RNA and so a DNA translocase would have immense trouble utilizing an identical mechanism; the stiffer dsDNA substrate in a DNA translocase channel would mean that each single protein contact would have to move almost 30 Å to maintain contact with the DNA during a full catalytic cycle around the hexameric ring. This amount of movement of a contact point within a tight hexameric structure seems unlikely. Further, it would be energetically unfeasible to compress the double helix greatly yet still produce power from ATP hydrolysis. Therefore, an intermediate or “limited-escort” model was proposed for FtsK whereby three adjacent monomers in a ring contact dsDNA simultaneously. This could allow for one inactive subunit to be skipped if the monomers on either side of it are proficient for translocation [23,175]. It would also imply that two adjacent mutant subunits would produce an inactive hexamer [175]. Both the inchworm model and the partial escort model are consistent with a mechanism that is largely that of a revolution motor. Both models propose that dsDNA touches the internal surface of

Biological Nanomotors: Linear, Rotation, or Revolution Mechanism

29

the hexameric ring and that the contact point between protein and DNA revolves around the inner surface of the protein multimer with minimal rotation [12,23,82]. Both models proposed that a defined number of bases would be moved per ATP hydrolyzed; if each subunit contacts the DNA substrate in an identical fashion around the ring, then the DNA must be moved by a defined length at each step to maintain these identical interactions. A slight twisting of the DNA at each step is then necessary to maintain the identity of the contacts with the dsDNA around the ring to maintain the register between DNA and protein; the angle between adjacent subunit active sites in a hexamer is 360°/6 or 60°, whereas the angle between adjacent phosphates, around the dsDNA axis, is 360°/10.5 (about 34°). If precisely 2 bp are translocated per subunit, then there is a requirement to twist the DNA an extra 8° to maintain the identity of each protein-DNA contact. DNA within cells is negatively supercoiled, with a supercoiling density in E. coli of around −0.05. Taking this into account, the amount of twisting required for each FtsK power step is reduced to around 5° per 2 bp translocated, which corresponds to one supercoil induced for every 144 bp translocated. This theoretical value is almost identical to the value of one supercoil per 150 bp observed using single-molecule experiments [174]. If this is the case, small over-winding of the DNA ahead of a translocating FtsK will produce positive supercoiling ahead of the protein. On the other hand, in the cell, this might be removed either by the action of DNA gyrase or by the occasional slipping of the motor to release the torsional tension in the DNA. With the close fit of the DNA into the central channel of FtsK, movement of the DNA relative to the protein would not require large amounts of rotation. The point(s) of contact between FtsK and DNA and the concomitant wave of ATP binding and hydrolysis would move in a counterclockwise manner as viewed from the DNA entry side if the hexamer, revolving around the inner surface of the motor protein, but with very little deviation of the DNA helix away from the helical axis. This could potentially aid in the efficiency and speed of translocation by FtsK and related proteins. Complicated motors are available with multiple functional modules. In eukaryotic cells, many motors such as those involved in homologous recombination, DNA repair, Holliday junction resolution, nuclear membrane-embedded pore, or other membrane transporters are composed of multiple functional modules present as hexamer of other oligomers. For example, Holiday Junction resolvase RuvB complex is made up of two hexamers [178,179]; DNA double-strand break (DSB) involves exonuclease, helicase, topoisomerase, DNA polymerase, and molecular zipper (180). Although the hexameric ATPase could be part of these motors, the involvement of more than one functional modules makes the classification challenging. Chromosome or genomic DNA either in eukaryotic cells or in bacteria, as a very long string, would cause coiling or tangling with rotation of such long double helix, and resolution of the coiling requires the consumption of huge energy. Thus, it is expected that some of the functional complexes within these complicated motors might belong to the revolution motor, since the use of the revolution mechanism will avoid dsDNA coiling and tangling as described in the above sections.

1.4.4

MeCHanisM oF lineaR MotoRs

Linear motors are highly efficient with estimates of 50%–60% of the energy from ATP hydrolysis utilized for mechanical motion [4,5,181]. In myosin, the actin filament is proposed to accelerate the structural changes in the force-generating element, which couples the mechanical and chemical cycles. The lever arm consists of the light chain-binding region. Variability in the length of the light chain-binding region in different myosin isoforms helped to prove the hypothesis that this region functions as a lever arm [6]. In kinesin, the major difference is that the ATP-binding step is associated with force generation, while hydrolysis step occurs with kinesin bound to the microtubule [18]. The neck linker or coiled-coil stalk has been demonstrated to be critical for movement. In dynein, the coiled coil of the stalk that connects the AAA ring and the microtubule-binding domain is known to change conformation in a nucleotide-dependent manner and function as a force-generating element [19].

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FIGURE 1.23 Crystal structure showing the different subdomains of myosin along with the actin, nucleotide-binding, and lever-arm regions. PDB code: 1W7J.

Conserved catalytic cycle of myosins. The modified Lymn-Taylor cycle provides the minimal framework for explaining the conserved properties of the actomyosin ATPase cycle [182]. The motor domain of myosin contains the sites for ATP binding (nucleotide-binding pocket—NBP) and actin binding (actin-binding cleft—cleft) [183]. These sites are coordinated with the reciprocal movement of the lever-arm region during the recovery and power stroke states of the ATPase cycle (Figure 1.11). In the absence of any nucleotide, myosin binds to actin tightly and forms a strongly bound complex. ATP binding to myosin causes the cross-bridge to detach from actin and enter the weak binding states. During the detached states, ATP is hydrolyzed by myosin, and the lever-arm region of myosin primes itself into a pre-powerstroke state (Recovery Stroke, Figure 1.11) [184]. Thereafter, myosin complexed with the hydrolysis products rebinds to actin in a weak binding state. Actin binding activates the release of phosphate from the active site, which is a key step in coupling the mechanical and chemical cycles, and then the release of ADP occurs when myosin is strongly bound to actin. During the actomyosin-bound state, myosin pulls on the actin filament performing mechanical work, which is produced by the swing of the lever arm (Powerstroke, Figure 1.11). Nucleotide-binding region. The coordination of ATP within the nucleotide-binding pocket, cleavage of its phosphoanhydride bond, and the sequential release of products governs the mechanical cycle of myosins [185–187]. Small structural changes in the conserved regions of the nucleotidebinding pocket are communicated to the actin-binding and lever-arm regions (Figure 1.23). Switch I and Switch II coordinate the sequential release of products and transmit information from the NBP to the actin-binding and lever-arm regions [188]. Switch II forms a salt bridge with Switch I and also interacts with the γ-phosphate of ATP [189,190]. The P-loop is also involved in the coordination of the α and β phosphates of ATP. Additionally, magnesium (Mg) is coordinated to the oxygen on the β and γ phosphates and makes a direct or water-mediated contact with residues of Switch I (Figure 1.24). The switch elements undergo a conformational change to a “closed” state upon binding of ATP, which leads to a twisting of a seven-stranded β sheet (transducer), resulting in the opening of the actin-binding cleft [191]. Moreover, the twist of the transducer region also translates towards the C-terminal, lever-arm region via a highly conserved structural element called the relay helix (Figure 1.24). The relay helix-communication pathway induces the recovery stroke

Biological Nanomotors: Linear, Rotation, or Revolution Mechanism

31

FIGURE 1.24 Crystal structure of myosin showing the key structural elements involved in the coordination of ATP and the energy transduction mechanism as discussed in the text. PDB code: 1W7J.

and formation of the pre-powerstroke state of the lever arm. The actin-binding cleft is a deep cleft between the upper (U50) and lower (L50) 50-kDa subdomains in the motor domain (Figure 1.23). The binding of myosin to actin is proposed to cause the movement of Switch I, which induces a loss of Mg coordination, reducing its affinity and its eventual release [192]. These and other rearrangements to the active site result in an isomerization to the weak ADP state of the pocket and eventual release of ADP. Actin-binding region. The open-closed transition of Switch I may be coupled to the closed-open equilibrium of the actin-binding cleft. A 32-amino acid-long alpha helix that traverses the upper 50-kDa domain of myosins, called the HO helix, and a related HG/HH helix have been demonstrated by molecular modeling studies to be strongly coupled during the open-close transition of the cleft [193] (Figure 1.24). Conformational changes in the HO helix during the myosin ATPase cycle correlate with ATP-induced dissociation and attachment to actin as demonstrated by intrinsic tryptophan fluorescence [7]. The relay helix near the lower 50-kDa domain is a 4.7-nm-long α-helix that has been well documented to be an essential feature of the force-generating region of myosin [184,194,195]. It connects the nucleotide-binding site to the lever-arm region and goes from a kinked to a straight conformation during the formation of the pre-powerstroke state [184,194,195]. The HO helix and the relay helix are connected via the Switch II loop. Lever-arm region. The lever-arm movement during the power and recovery stroke stages of the catalytic cycle has been probed by a number of studies, including intrinsic tryptophan fluorescence [196,197], polarization [8,9], electron paramagnetic resonance [10], and more recently FRET [195]. There is a still a controversy about the timing of the power stroke in relationship to the product release steps of the ATPase cycle. Examining the kinetics of relay helix straightening by FRET during actin-activate phosphate release suggests that the power stroke occurs after actin binding and before phosphate release [195]. A study that directly measured the movement of the lever arm by FRET in myosin V demonstrated two phases of the power stroke: a rapid phase that occurs before phosphate release and a slower phase that occurs after phosphate release but before ADP release

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[11]. However, a recent crystal structure of myosin VI provides evidence that phosphate release gates the movement of lever associated with the power stroke [12]. There is good agreement on the details of movement of the lever arm during the recovery stroke. A FRET study reported that the reverse movement of the relay helix from a straight to a kinked conformation is associated with the reversal of the powerstroke or the recovery stroke [195]. Other studies are in agreement that the straight-to-kinked transition of the relay helix occurs after ATP binding and before hydrolysis [184,194]. The structural mechanism of dynein-based motility is not well understood because of the lack of high-resolution structure. However, several recent reports have used a combination of EM and X-ray crystallography to shed light in the details of the key structural changes that drive microtubule binding and force generation [198]. In the presence of an ATP analog, there is closure of the motor ring, which drives the movement of the linker domain, a structural element proposed to be crucial for force generation. The coiled-coil helix of the stalk domain also slides in a nucleotide-dependent manner, which alters the affinity for microtubule. Thus, dynein is similar to myosin in that the ATP binding and hydrolysis prime the motor for force generation, while binding to the track triggers the force-generating structural change.

1.4.5

MeCHanisM in ContRol sequential and CooRdination aMong CHannel subunits

Nucleic acid translocation and duplex unwinding by helicases are coupled to the NTP-binding and hydrolysis cycle. In nucleic acid translocation, there are six NTP-binding sites, as well as at least six nucleic acid-binding sites in the hexamer. Individual subunits of a hexameric helicase may switch between several DNA-binding states. The binding of NTP to a hydrolysis site, near the inner surface of the channel, induces a conformational change that exposes one pair of negatively and positively charged regions per nucleotide-hydrolysis site. These charged regions are not of equal size, are oriented at an angle to the circumferential meridian, and are not constant, appearing and disappearing with the binding and hydrolysis of NTP, respectively. The field of these charged regions can sequentially interact with the closest negatively charged DNA phosphate, which gives an “electrostatic push” in the direction of the charged pair axis. The combined effect of the charged pairs produces a sustained torsional and axial thrust. Sequential action of phi29 DNA-packaging motor was originally reported by Chen and Guo [199] and subsequently confirmed by Bustamante and coworkers [116]. Hill constant determination and binomial distribution of inhibition assay have led to the conclusion that ATPase subunits work sequentially and cooperatively [12,49]. This action enables the motor to work continuously without interruption, despite some observable pauses. Motor subunits work cooperatively, as deduced from the observation of the binding of dsDNA to only one gp16 subunit at a time. ATPase activity has been analyzed by studying the effects of introducing mutant subunits on the oligomerization of gp16 [114,115]. In the absence of dsDNA, increasing the ratio of gp16 Walker B mutants failed to produce a significant effect on ATPase activity. However, when saturating amounts of dsDNA were added, a strong negative cooperative effect was produced. The profile mostly overlapped with a model where one single inactive subunit was able to inactivate the whole oligomer. The model was calculated from the binomial distribution inhibition assay [114,115]. These results suggest that in the presence of dsDNA, a rearrangement occurs within the subunits of gp16 that enables them to communicate with each other and to “sense” the nucleotide state of the reciprocal subunit. DNA translocation occurs, while the other ATPase subunits are in a type of “stalled” or “loaded” state. This represents an extremely high level of coordination between proteins with their DNA substrate, perhaps the most efficient process of coupling energy production via ATP hydrolysis with DNA translocation of all viral motors known so far. An effective mechanism of coordination is apparent between gp16 and dsDNA regulated by the ATP hydrolysis cycles. The cooperativity and sequential actions among hexameric ATPase subunits [116,199] promote the revolution of dsDNA along the ATPase channel.

Biological Nanomotors: Linear, Rotation, or Revolution Mechanism

1.5

33

POTENTIAL MOTOR APPLICATIONS

The discoveries discussed in this review offer a series of possible answers to the puzzles in nanotechnology that may lead to inventions in the motion world. The riding system along one string of dsDNA provides evidence of a motion system for cargo transportation at the nanoscale, and a tool for studying force generation mechanisms in a moving world. The revolution mechanism itself offers a prototype or a hint for the design of new motors involving forward motion via a tract, such as that used by roller coasters, trolley cars, or launcher for flying objects. Nature has evolved a clever machine that translocates DNA double helix to avoid difficulties associated with rotation, e.g., DNA supercoiling. These nanomachines can also be applied for the construction of sophisticated nanodevices, including molecular sensors [200–202], bioreactors [203], chips, DNA-sequencing apparatus [166,200,204,205], or other electronic and optical devices [206,207]. Nanopore-based analysis has a wide range of incredibly versatile applications, including the sensing of small molecules such as ions, nucleotides, and drugs, as well as larger polymers such as PEG, nucleic acid, and polypeptides [128,200–202,208]. In addition, these multi-subunit biocomplexes have a potential to serve as drug targets for therapeutics, as well as diagnostic applications [201,202,209,210], especially the biomotors with an advantage of a high stoichiometry, which may provide clues to solve the drug-resistant issue [211,212].

1.6

CONCLUDING REMARK AND PERSPECTIVES

Nanobiomotors are tiny machines that utilize a primary energy source into mechanical works. They are crucial to the sustenance of living systems, since they provide for the most forms of biological motion; help direct cellular components to proper destinations, package DNA, contract muscles; and perform a variety of other functions. Biomotors exhibit a diversity of complex structures. Most have the same basic components, including a mechanical frame (usually composed of proteins) with both moving and static parts, powered by an energy supply. This energy is typically derived from the hydrolysis of ATP, which leads to conformational changes in the motor protein, resulting in movement, but other motors use energy produced from ion gradients. These motors are typically divided into categories based on the type of motion displayed: the most well-studied motors are categorized tentatively into linear motors, rotary motors, and nucleic acid-translocating motors. Biomotors were once classified into two categories: linear motor and rotation motor. For decades, the viral DNApackaging motor has been popularly believed to be a fivefold rotation motor. Recently, a third type of biomotor with revolution mechanism without rotation has been discovered. By analogy, rotation resembles the Earth rotating on its axis in a complete cycle every 24 hours, while revolution resembles the Earth revolving around the Sun one circle per 365 days (see animations http://nanobio.uky. edu/movie.html). The action of revolution that enables a motor free of coiling and torque has solved many puzzles and debates that have occurred throughout the history of viral DNA-packaging motor studies. It also settles the discrepancies concerning the structure, stoichiometry, and functioning of DNA translocation motors. This review uses bacteriophages Phi29, HK97, SPP1, P22, T4, T7 as well as bacterial DNA translocase FtsK and SpoIIIE as examples to elucidate the puzzles. These motors all use a hexameric ATPase to revolve around the dsDNA sequentially. ATP binding induces conformational change and possibly an entropy alteration in ATPase to a high affinity towards dsDNA; but ATP hydrolysis triggers another entropic and conformational change in ATPase to a low affinity for DNA, by which dsDNA is pushed towards an adjacent ATPase subunit. The rotation and revolution mechanisms can be distinguished by the size of channel: the channels of rotation motors are equal to or smaller than 2 nm, whereas channels of revolution motors are larger than 3 nm. Rotation motors use parallel threads to operate with a right-handed channel, while revolution motors use a left-handed channel to drive the right-handed DNA in an anti-parallel arrangement. Coordination of several vector factors in the same direction makes viral DNA-packaging motors unusually powerful and effective. Revolution mechanism avoids DNA coiling in translocating the lengthy genomic dsDNA helix.

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ACKNOWLEDGMENTS The authors thank Dr. Darshan Trivedi, Stanford University, for generating Figures 1.11, 1.23, 1.24 and for critical comments; Dr. W. E. Moerner, Stanford University; Dr. Imre Derenyi, Eötvös University, Hungary; Dr. P. Satir, Yeshiva University; Dr. Konrad J., Böhm Institute of Molecular Biotechnology, Germany; Dr. Yoshie Harada, The Tokyo Metropolitan Institute of Medical Science; Dr. Eckhard Jankowsky, Case Western Reserve University; Dr. Ron Vale, University of California; Dr. Nadrian Seeman, New York University; Dr. Peter Prevelige Jr., University of Alabama; Dr. Kazuhiko Kinosita, Okazaki National Research Institutes; Dr. Christof Niemeyer, Dortmund University, Germany; Dr. Toshio Yanagida and Dr. E. Taniguchi, Osaka University, for critical reading of the manuscript and helpful discussions. The work was supported by NIH Grants R01-EB003730, R01-EB012135, R01-EB019036, and U01-CA151648 to PG; and an AHA Grant to CMY. The content is solely the responsibility of the authors and does not necessarily represent the official views of NIH. Funding to Peixuan Guo’s Endowed Chair in Nanobiotechnology position is from the William Fairish Endowment Fund.

COMPETING INTERESTS PG is a co-founder of Biomotor and RNA Nanotech Development Co. Ltd.

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Classifications and Typical Examples of Biomotors Peixuan Guo and Zhengyi Zhao University of Kentucky

CONTENTS 2.1

Typical Revolving Motors....................................................................................................... 43 2.1.1 DNA Packaging Motor of Double-Stranded DNA Bacteriophages ...........................44 2.1.2 DNA Packaging Motor of Eukaryotic dsDNA Viruses.............................................. 45 2.1.3 dsDNA Translocases FtsK/SpoIIIE Superfamily ....................................................... 47 2.2 Typical Rotary Motors ............................................................................................................ 47 2.2.1 FoF1 Complex .............................................................................................................. 48 2.2.2 DNA Helicase ............................................................................................................. 48 2.2.3 Bacterial Flagella ........................................................................................................ 48 2.3 Typical Linear Motors ............................................................................................................ 48 References ........................................................................................................................................ 49

2.1

TYPICAL REVOLVING MOTORS

Experimental evidence shows that the non-rotational properties of these biomotors include the following: (i) the motor is still active in packaging with the connectors fused to the procapsid protein, making rotation impossible (Baumann et al., 2006); (ii) results from single-molecule force spectroscopic studies combined with polarization spectroscopic studies showed no signals of connector rotation (Hugel et al., 2007); (iii) no rotation of the bead clusters, which were tethered to the DNA ends, was observed during active packaging (Chang et al., 2008). DNA was found to twist by as little as 1.5 degree per base pair translocated (Liu et al., 2014), confirming a non-rotational mechanism since one helical turn of dsDNA is ~10.5 bases and 1.5°/bp × 10.5 bp/turn = 15.7° is far below 365° per complete helical turn. The way that revolving motors perform work is somewhat similar but distinct to rotating motors in that they have multiple parts, one of which engages in circular motion; however, this movement is not akin to how the Earth spins on its own axis, but to how the Earth revolves around the Sun (Figure 2.1) (see animations http://rnanano.osu.edu/movie.html). This characteristic is widespread in a large variety of motors, including dsDNA viruses, dsDNA bacteriophages, and bacteria. Many members of this family are DNA packaging motors or bacterial dsDNA translocases. While there are some structural differences between these biomotors, they share the same basic components, namely, a hexameric ATPase ring and a central channel through which DNA passes. The role of this powerful motor in dsDNA viruses is to inject a single piece of DNA into a procapsid protein shell. Bacteriophages are viruses consisting of an infectious tailpiece and a procapsid (head) made of protein. Viruses themselves cannot reproduce on their own; they do so via the genetic material (DNA or RNA) that they hijack. Because the DNA genome is very long when stretched out, the motor must exert a tremendous amount of force in order to cram the material into the viral capsid; when released, this pressure forces the DNA into an infected host cell. The injected DNA instructs the bacterium to build proteins, automatically assembling into new capsids. It kills off the bacterium, then attaches itself to other bacteria, and repeats the process of infection until every cell has been killed. DOI: 10.1201/9780429203367-3

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Biomotors and their Nanobiotechnology Applications

FIGURE 2.1 Illustration of different categories of motors. Linear motors are like people walking (PDB code 3KIN); Rotation motors are like a wheel and like Earth rotating on its own axis; Revolution motors resemble Earth revolving around the sun without self-rotation.

Other revolving viral DNA packaging motors include the T4, said to be twice as powerful as an automobile engine, the herpes virus, and FtsK (filamenting temperature-sensitive mutant K), which comprises a family of motor proteins. Located within eukaryotic cells, FtsK is responsible for coordinating the accurate copying of the entire complement of genetic material and the late stages of chromosome segregation with cell division. It is especially important for cell division in Escherichia coli.

2.1.1

dna paCKaging MotoR oF double-stRanded dna baCteRiopHages

The revolving mechanism was first proposed in 2013 in the study of the well-studied phi29 dsDNA packaging motor (Schwartz et al., 2013; Zhao et al., 2013). Inspired by the findings in the phi29 biomotor, studies in the other dsDNA motors have been carried out. It was confirmed that the revolving mechanism is a common feature shared by all dsDNA packaging motors, including SPP1, P22, T7, and the HK97 family phage, evidenced by the results from both crystal structural and biochemical studies. During replication, the phi29 bacteriophage translocates its genomic DNA into procapsids (Guo & Lee, 2007; Rao & Feiss, 2008; Zhang et al., 2012; Serwer, 2010), packing DNA against an ever-increasing internal pressure. To overcome this entropically unfavorable process, an energetically favorable motion is required to be coupled to the process of packaging (Guo et al., 1987b; Chemla et al., 2005; Hwang et al., 1996; Sabanayagam et al., 2007). Viral dsDNA packaging motors consist of a dodecameric center channel (Jimenez et al., 1986; Guasch et al., 2002), a hexameric packaging RNA ring (Guo et al., 1987a; Guo et al., 1998; Shu et al., 2007; Zhang et al., 2013), and an ATPase hexameric ring (Figure 2.2). Crystal structural analysis of all the motor channels of SPP1 (Lebedev et al., 2007), T7 (Agirrezabala et al., 2005), HK97 (Juhala et al., 2000), P22 (Olia et al., 2011), and phi29 (Guasch et al., 2002) revealed the existence of a common anti-chiral arrangement between their channel subunits and the dsDNA helices. It has also been found that in many viruses; dsDNA is naturally spooled inside the viral capsid after packaging, free from rotation tangles (Lander et al., 2006; Molineux & Panja, 2013; Petrov & Harvey, 2008; Jiang et al., 2006).

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FIGURE 2.2 Depiction of the structure and function of the phi29 DNA-packaging motor. (upper panel) Side view of phi29 dsDNA-packaging motor (left) and top view of phi29 connector (right). (lower left) Hexameric pRNA generated from crystal structures of its 3WJ core and AFM images of loop-extended hexameric pRNA. (lower right) DNA revolving inside the connector channel by contact with each connector subunit in a 30° transition step for each contact.

In phi29, a toroid of dsDNA has been shown by Cryogenic electron microscopy (cryo-EM) around the portal region (Figure 2.3) (Tang et al., 2008; Sherratt et al., 2010), which is consistent with the revolving mechanism since such toroidal structural images may result from the accumulation of individual revolving DNA processed by Cryo-EM. A compression mechanism found in the T4 DNA packaging motor (Dixit et al., 2012; Ray et al., 2010) also agrees with the revolving mechanism and disagrees with the pentameric gp17 model of T4.

2.1.2

dna paCKaging MotoR oF euKaRyotiC dsdna viRuses

Adenoviruses (AdV) are a group of well-studied dsDNA viruses that infect eukaryotic cells of vertebrates, including humans. AdV packages the genome with two subunits of the terminal proteins into capsids that contain hexon, penton, and fiber (La et al., 2003). Iva2 and L1 52/55K proteins are AdV packaging proteins. It has been reported that higher-order IVa2-containing complexes formed on adjacent packaging repeats are required for packaging activity (Ostapchuk et al., 2005). Quantitative

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FIGURE 2.3 Examples of spooling of DNA within capsids of phages to support the revolution mechanism. (upper left) Bacteriophage phi29. (lower left) Bacteriophage P22. (right panel) Bacteriophage T7

mass spectrometry, metabolic labeling, and Western blot revealed that there are approximately six to eight IVa2 molecules in each particle (Christensen et al., 2008). The main motor protein Iva2 in AdV is multifunctional. It assists in the assembly of the capsid and activates late transcription. Comparing the IVa2 protein sequences with ATPase from different species revealed conserved Walker A and B motifs associated with binding and hydrolysis of ATP (Ostapchuk & Hearing, 2008). Similar to the ATPases in bacteriophage packaging motors, the multimeric ATPase IVa2 motor protein complex also works through a sequential action to provide energy for the packaging of DNA into their capsids. IVa2 also interacts with a viral L4-22K protein, which has been shown to be involved in genome encapsidation (Ostapchuk et al., 2006; Ewing et al., 2007; Tyler et al., 2007). IVa2 mutants were defective in DNA packaging and resulted in the accumulation of empty capsids similar to the procapsid of dsDNA bacteriophages (Christensen et al., 2008) (see http://www.ncbi. nlm.nih.gov/pmc/articles/PMC4404914/ - B6.) Herpes simplex viruses (HSV) package their dsDNA genome into a preformed protein shell using terminase (Raoult et al., 2004), which contains a large subunit pUL15 and a small subunit pUL28 (Koslowski et al., 1999). pUL15 cleaves concatemeric viral DNA during packaging initiation and completion cycles and functions as an ATPase, providing energy to the packaging process. X-ray structure of the C-terminal domain of pUL15 showed a homo-trimer structure (Raoult et al., 2004). The structure of the C-terminal domain of pUL15 resembles those of bacteriophage terminases, RNase H, integrases, DNA polymerases, and topoisomerases, with an active site clustered with acidic residues. The DNA-binding surface is surrounded by flexible loops, indicating that they adopt conformational changes upon DNA binding (Selvarajan et al., 2013). These conformational changes are similar to the sequential action of ATPase gp16 observed in phi29 DNA packaging motors, which provides energy to support the one-way traffic of genome into procapsids. Mimivirus, megavirus, pandoravirus, and pithovirus (Philippe et al., 2013; Arslan et al., 2011; La et al., 2003) all belong to the nucleocytoplasmic large DNA viruses (NCLDV) superfamily, and infect a wide range of eukaryotes (Chelikani et al., 2014b; Ghedin & Fraser, 2005). Nine genes that are shared by all NCLDV families have been identified to encode for DNA polymerase: a capsid protein, 3 helicases, a virion packaging ATPase, a thiol oxidoreductase, a protein kinase, and a transcription factor (Raoult et al., 2007). Mimiviruses package their 1.2-Mbp dsDNA genome into preformed procapsids through a non-vertex portal (Zauberman et al., 2008) driven by the vaccinia virus A32-type virion packaging ATPase (Monier et al., 2008). It has been shown that the structure and function of their DNA packaging motors are homologous to the FtsK DNA translocase

Classifications and Typical Examples of Biomotors

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(Chelikani et al., 2014b; Chelikani et al., 2014a). The genome packaging motors of NCLDVs interact with other genome packaging components such as recombinase and type II topoisomerase similar to prokaryotic FtsK DNA translocase (Chelikani et al., 2014b; Chelikani et al., 2014a). Poxviruses, another member of NCLDVs, are large, brick-shaped dsDNA viruses that replicate in the cytoplasm of infected cells. The two DNA strands of the genome are connected at the ends through hairpin termini (Moss, 1985). Poxvirus ATPase is coded by the A32 gene; comparative sequence analysis revealed a highly conserved N-terminal region with five motifs among all poxviruses, including ATPase featuring Walker A and Walker B motifs, A32L specific motifs III and IV, and a novel motif-V. The secondary structure predictions of N-terminus of A32 ATPase protein are homologous to those of the FtsK DNA translocase (Yogisharadhya et al., 2012). Baculoviruses are large viruses with circular dsDNA genomes that infect invertebrates (Huang et al., 2011). These viruses encode a motor protein (Ac66/ORF66), which is believed to be a key component of genome packaging. This protein has shown similarity with eukaryotic structural and chromosome maintenance protein (SMC) (Hoffmann & Thio, 2007; Huang et al., 2011; Deng et al., 2007).

2.1.3

dsdna

tRansloCases FtsK/spoiiie supeRFaMily

While there are different types of FtsK, they all share the same three components: an N-terminal domain with four transmembrane segments that anchor the motor to the cytoplasmic membrane, a C-terminal domain consisting of about 512 amino acids, and a long, winged helix linker that connects the N- and C-terminal domains and is rich in amino acids glutamine and proline. While the N-terminal domain plays a major role in cell division, the C-terminal domain functions in chromosome segregation and DNA translocation. The latter is divided into three segments: α, β, and γ. The first two parts are the motor portion of FtsK, and they assemble into a hexameric ring with a central channel through which dsDNA passes. The γ subdomain provides for directionality of the DNA. It has been found that the C-terminal domain of FtsK uses the γ subdomain to bind to DNA at sites of 8-basepair sequences called KOPS, which direct the rapid translocation of DNA. As with many other biomotors, it is the hydrolysis of ATP that powers this process, allowing FtsK to translocate up to 5,000 base pairs of DNA per second. FtsK is often grouped in the same superfamily as SpoIIIE, which is also involved in DNA conjugation, segregation, and translocation as well as protein transport. The FtsK/SpoIIIE family of proteins belongs to the ASCE (additional strand conserved E) superfamily, which are hexameric dsDNA translocases found in many bacterial species. Within the ASCE class, the FtsK/HerA clade is present throughout bacteria and archaea. The large FtsK-HerA family (Iyer et al., 2004) also contains the motor proteins of various conjugative plasmids and transposons such as the single-strand translocase protein TrwB (Gomis-Ruth & Coll, 2001; Gomis-Ruth et al., 2001).

2.2

TYPICAL ROTARY MOTORS

Rotation biomotors share a similar characteristic of movement: as the name would suggest, they perform work via rotation, in the same way that the Earth spins on its own axis. The category of rotation motors comprises a large number of classes, including bacterial flagella motors, helicases, and the FoF1-ATP synthase family of proteins (or FoF1-ATPase for short) (Junge et al., 1997; DeRosier, 1998; Enemark & Joshua-Tor, 2006). Bacterial flagella motors lie at the base of flagellar filaments and allow a bacterium to “swim.” FoF1-ATPase plays an indispensable role by converting proton gradients into ATP, which powers nearly all cellular processes. Helicases separate the two strands of the double-stranded DNA helix into single strands, making it possible for them to be replicated, in addition to many other functions.

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2.2.1

Biomotors and their Nanobiotechnology Applications

FoF1 CoMplex

FoF1 ATP synthase, as an ATP generator, is located in the inner mitochondrial membranes, the thylakoid membranes of chloroplasts, and the plasma membranes of bacteria (Yoshida et al., 2003; Okuno et al., 2011). Its two components, rotary motors Fo and F1, are connected by a common rotor axle and peripheral stalk. This typical rotary molecular energy converter interconverts proton motive force (pmf) and energy of ATP through the rotation of its subunit complex (Noji & Yoshida, 2001).

2.2.2

dna HeliCase

DNA helicases are a primary class of enzymes in genome maintenance. They serve a vital role in cell biology by breaking the hydrogen bonds between the double helix base pairing of doublestranded DNA, allowing the individual strands to be copied, repaired, and recombined. Once they are copied, new strands of DNA can be synthesized by DNA polymerase. Helicases are so ubiquitous that mutations or dysfunctions can cause a number of seemingly unrelated diseases and conditions, including cancer, immunodeficiency, and premature aging. The energy source for these biomotors is derived from the hydrolysis of NTP, typically ATP. Upon binding with DNA, the motor’s structure is reinforced and NTPase activity is stimulated. Most helicases are hexameric, though some can be dimeric or monomeric. Much like the F1 motor of FoF1-ATPase, the hexamer forms a ring shape with a central channel. What moves through this channel is a single strand of dsDNA, where the DNA strand serves as the stator and the helicase the rotor of the typical rotating motor. The bond between the strand and the channel is very close, and the helicase easily slides over the strand because both are right-handed and their threads parallel, in the same way that a screw passes through a bolt when it is turned. Nucleic acid helicases can be classified through numerous ways. On the basis of amino acid sequence comparisons, helicases are divided into three large superfamilies (F1, F2, and F3) and two smaller families (F4 and F5). DNA helicases and RNA helicases can both be found in each of these helicase superfamilies except for F6 (Jankowsky & Fairman-Williams, 2010). F5 contains a bacterial Rho factor involved in transcription termination regulation, and F6 contains a structure that functions similar to the Rho factor but is different in structure (Jankowsky, 2010). The cores of F1 and F2 enzymes are structurally homologous and contain similar sets of seven helicase signature motifs.

2.2.3

baCteRial Flagella

Flagella, with a still helical-shaped structure, are the major force generators for most motile bacteria. Each flagella filament has an individual rotation motor at its base (DeRosier, 1998; Berry, 2001), which can rotate clockwise to generate forward movement, or counterclockwise for tumbling or direction alternation. The flagellar motor is powered by proton motive force with ATP as an energy source. A number of models have been proposed to explain the mechanism of flagellar motor function (Khan & Berg, 1983; Berg & Turner, 1993; Atsumi, 2001; Walz & Caplan, 2000; Thomas et al., 1999; Berry & Berg, 1999; Elston & Oster, 1997; Berry, 1993; Lauger, 1988), for example, the “electrostatic proton turbine” model (Elston & Oster, 1997; Berry, 1993; Lauger, 1988) and the “turnstile” model (Khan & Berg, 1983; Berg & Turner, 1993). Since many excellent updated reviews are available, readers are encouraged to refer to Guo et al., 2014; Guo et al., 2013; Stevenson et al., 2015 Dutcher, 2014; Fisch & Dupuis-Williams, 2011; Van et al., 2011; Inaba, 2011; Minamino et al., 2008.

2.3

TYPICAL LINEAR MOTORS

Linear motors were the first described motor proteins. Unlike the rotation mechanism of the previous class of motors, they advance unidirectionally along a cytoskeletal track. This category includes three main types of motors – myosin, kinesin, and dynein – each of which counts a large number of

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subtypes. Like the majority of other motors, they depend on ATP to produce mechanical work. They are responsible for most of the movements in eukaryotic cells. Myosin, for example, plays a primary role in muscle contraction as well as a variety of other intracellular functions. Kinesin is involved in several activities, including meiosis, mitosis, and intracellular transport of cellular cargo.

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Virol J 5, 12. Moss, B., 1985. Replication of poxviruses. In: B. N. Fields, et al. (Eds.), Virology. Raven Press, New York, pp. 685–703. Okuno, D., Iino, R., Noji, H., 2011. Rotation and structure of FoF1-ATP synthase. J Biochem 149, 655–664. Olia, A. S., Prevelige, P. E., Johnson, J. E., Cingolani, G., 2011. Three-dimensional structure of a viral genomedelivery portal vertex. Nat Struct Mol Biol 18, 597–603. Ostapchuk, P., Anderson, M. E., Chandrasekhar, S., Hearing, P., 2006. The L4 22-kilodalton protein plays a role in packaging of the adenovirus genome. J Virol 80, 6973–6981. Ostapchuk, P., Hearing, P., 2008. Adenovirus IVa2 protein binds ATP. J Virol 82, 10290–10294. Ostapchuk, P., Yang, J., Auffarth, E., Hearing, P., 2005. Functional interaction of the adenovirus IVa2 protein with adenovirus type 5 packaging sequences. J Virol 79, 2831–2838. Petrov, A. S., Harvey, S. C., 2008. Packaging double-helical DNA into viral capsids: structures, forces, and energetics. Biophys J 95, 497–502. Philippe, N., Legendre, M., Doutre, G., Coute, Y., Poirot, O., Lescot, M., Arslan, D., Seltzer, V., Bertaux, L., Bruley, C., Garin, J., Claverie, J. M., Abergel, C., 2013. Pandoraviruses: amoeba viruses with genomes up to 2.5 Mb reaching that of parasitic eukaryotes. Science 341, 281–286. Rafferty, J. B., Sedelnikova, S. E., Hargreaves, D., Artymiuk, P. J., Baker, P. J., Sharples, G. J., Mahdi, A. A., Lloyd, R. G., Rice, D. W., 1996. Crystal structure of DNA recombination protein RuvA and a model for its binding to the Holliday junction. Science 274, 415–421. Rao, V. B., Feiss, M., 2008. The Bacteriophage DNA Packaging Motor. Annu Rev Genet 42, 647–681. Raoult, D., Audic, S., Robert, C., Abergel, C., Renesto, P., Ogata, H., La, S. B., Suzan, M., Claverie, J. M., 2004. The 1.2-megabase genome sequence of Mimivirus. Science 306, 1344–1350. Raoult, D., La, S. B., Birtles, R., 2007. The discovery and characterization of Mimivirus, the largest known virus and putative pneumonia agent. Clin Infect Dis 45, 95–102. Ray, K., Sabanayagam, C. R., Lakowicz, J. R., Black, L. W., 2010. DNA crunching by a viral packaging motor: compression of a procapsid-portal stalled Y-DNA substrate. Virology 398, 224–232. Sabanayagam, C. R., Oram, M., Lakowicz, J. R., Black, L. W., 2007. Viral DNA packaging studied by fluorescence correlation spectroscopy. Biophys J 93, L17–L19. Schwartz, C., De Donatis, G. M., Zhang, H., Fang, H., Guo, P., 2013. Revolution rather than rotation of AAA+ hexameric phi29 nanomotor for viral dsDNA packaging without coiling. Virology 443, 28–39. Selvarajan, S. S., Zhao, H., Kamau, Y. N., Baines, J. D., Tang, L., 2013. The structure of the herpes simplex virus DNA-packaging terminase pUL15 nuclease domain suggests an evolutionary lineage among eukaryotic and prokaryotic viruses. J Virol 87, 7140–7148. Serwer, P., 2010. 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Van, G. N., Waksman, G., Remaut, H., 2011. Pili and flagella biology, structure, and biotechnological applications. Prog Mol Biol Transl Sci 103, 21–72. Walz, D., Caplan, S., 2000. An electrostatic mechanism closely reproducing observed behavior in the bacterial flagellar motor. Biophys J 78, 626–651. Wang, M. D., Schnitzer, M. J., Yin, H., Landick, R., Gelles, J., Block, S. M., 1998. Force and velocity measured for single molecules of RNA polymerase. Science 282, 902–907. Yogisharadhya, R., Bhanuprakash, V., Venkatesan, G., Balamurugan, V., Pandey, A. B., Shivachandra, S. B., 2012. Comparative sequence analysis of poxvirus A32 gene encoded ATPase protein and carboxyl terminal heterogeneity of Indian orf viruses. Vet Microbiol. 156, 72–80. Yoshida, M., Muneyuki, E., Hisabori, T., 2003. ATP synthase--a marvellous rotary engine of the cell. Nat Rev Mol Cell Biol 2(9), 669–677. Zauberman, N., Mutsafi, Y., Halevy, D. B., Shimoni, E., Klein, E., Xiao, C., Sun, S., Minsky, A., 2008. Distinct DNA exit and packaging portals in the virus Acanthamoeba polyphaga mimivirus. PLoS Biol 6, e114. Zhang, H., Endrizzi, J. A., Shu, Y., Haque, F., Sauter, C., Shlyakhtenko, L. S., Lyubchenko, Y., Guo, P., Chi, Y. I., 2013. Crystal structure of 3WJ core revealing divalent ion-promoted thermostability and assembly of the phi29 hexameric motor pRNA. RNA 19, 1226–1237. Zhang, H., Schwartz, C., De Donatis, G. M., Guo, P., 2012. “Push through one-way valve” mechanism of viral DNA packaging. Adv Virus Res 83, 415–465. Zhao, Z., Khisamutdinov, E., Schwartz, C., Guo, P., 2013. Mechanism of one-way traffic of hexameric phi29 DNA packaging motor with four electropositive relaying layers facilitating anti-parallel revolution. ACS Nano 7, 4082–4092.

3

Structure of Revolving Biomotors Peixuan Guo and Zhengyi Zhao University of Kentucky

CONTENTS 3.1 Hexameric Arrangement of Motor Components .................................................................... 53 3.2 dsDNA Translocases of the FtsK/SpoIIIE Superfamily......................................................... 53 References ........................................................................................................................................ 55

3.1

HEXAMERIC ARRANGEMENT OF MOTOR COMPONENTS

In 1978, structural studies of viruses (Caspar & Klug, 1962; Bazinet & King, 1985; Cardarelli et al., 2010; Valpuesta et al., 1992; Kochan et al., 1984; Doan & Dokland, 2007; Carrascosa et al., 1982; Agirrezabala et al., 2005) led to the popular fivefold/sixfold symmetric mismatch gearing mechanism (Hendrix, 1978). Twenty years later, the pRNA complex on the phi29 bacteriophage was first shown to be hexameric by Guo et al. (Guo et al., 1998; Zhang et al., 1998) (featured by Cell (Hendrix, 1998)). Despite some divergent models indicating this pRNA as a pentamer (Chistol et al., 2012; Yu et al., 2010; Morais et al., 2008), the hexameric configuration of pRNA was verified by various approaches, such as cryo-electron microscopy (Cryo-EM) (Ibarra et al., 2000), biochemical analysis (Guo et al., 1998; Zhang et al., 1998; Hendrix, 1998), single-molecule photobleaching step counting (Shu et al., 2007), electron microscopy (EM) (Moll & Guo, 2007; Xiao et al., 2008), and crystallization studies (Zhang et al., 2013). The photobleaching study of pRNA was carried out under the active packaging of the phi29 motor, excluding the possibility that the pRNA complex functions as a pentamer with one of its subunit falling off after packaging initiation (Morais et al., 2001; Simpson et al., 2000; Morais et al., 2008). Members of the additional strand conserved E (ASCE) superfamily mostly operate with a hexameric arrangement of its components (Iyer et al., 2004; Mueller-Cajar et al., 2011; Wang et al., 2011; Aker et al., 2007; Willows et al., 2004; Chen et al., 2002). Many revolving motors belong to the ASCE superfamily, including the FtsK hexamer and phi29 ATPase, which has been recently proven to adopt a hexamer configuration as its final oligomeric state (Schwartz et al., 2013) by virion assembly inhibition assays, binomial distribution analysis, qualitative DNA binding assays, capillary electrophoresis (CE) assays, and electrophoretic mobility shift assays (EMSA) (Chen et al., 1997; Trottier & Guo, 1997; Schwartz et al., 2013) (Figure 3.1).

3.2

dsDNA TRANSLOCASES OF THE FtsK/SpoIIIE SUPERFAMILY

The FtsK motor contains three functional components: one for DNA translocation, one for orientation control, and one for anchoring itself to the bacterial membrane (Figure 3.2) (Demarre et al., 2013). The N-terminal domain of FtsK consists of four transmembrane helices and interacts with FtsZ, thus leading to its localization and to the forming septum at mid-cell just prior to division (Yu et al., 1998; Dubarry et al., 2010). FtsK also interacts with a number of downstream cell division proteins to localize them to the divisome and allow cell division to progress (Dubarry et al., 2010). The N- and C-terminal domains are connected by a linker domain rich in proline and glutamine, DOI: 10.1201/9780429203367-4

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FIGURE 3.1 Stoichiometric assays showing the formation of the phi29 ATPase hexamer. (upper left) A native gel reveals six oligomeric states of the ATPase; the hexamer formation increases as the concentration of protein is increased. (lower left) Quantification by varying the [ATPase]/[DNA] molar ratio. The concentration of bound DNA plateaus at a molar ratio of 6:1. (right panel) A slab gel showing the binding of ATPase to dsDNA in a 6:1 ratio, imaged in GFP (upper panel) and Cy3 (lower panel) channels for ATPase and dsDNA, respectively.

FIGURE 3.2 Model of the FtsK motor loaded at a KOPS site. The N termini of the γ domains are located on one side of the complex, where they would connect to the motor domains of FtsK. This leads to loading of the motor to one side of the KOPS site so that the motor is pointing in a defined direction. This gives the motor its subsequent directional translocation (the arrow denotes the direction the motor would move along the DNA).

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FIGURE 3.3 The FtsK motor protein. (upper panel) E. coli FtsK protein domain structure. (middle panel) Two views of the hexameric FtsK motor protein structure. (lower panel) Structure of three γ domains bound to a KOPS DNA, seen along the DNA axis (left) and from the side (right).

which also aids in interactions to localize the cell division apparatus. The C-terminus consists of the DNA translocation motor ATPase, which drives DNA away from the forming septum and also participates in XerCD site-specific recombination at dif sites to resolve chromosome dimers. The C-terminal DNA translocation motor component of FtsK can be further subdivided into three domains, α, β, and γ (Massey et al., 2006). The α and β domains assemble into a hexameric ring-shaped channel, through which the dsDNA substrate is threaded (Massey et al., 2006), as revealed by EM. The rings formed by α and β are separated by a ~10Å cleft and connected by two strands. The γ subdomain acts as both a protein-protein domain and DNA binding domain. It activates Xer-mediated recombination at dif (Grainge et al., 2011) and recognizes and binds to the specific 8-bp chromosomal sequences (GGGNAGGG) (Figure 3.3) (Lowe et al., 2008) known as FtsK orienting polarized sequences (KOPS) (Bigot et al., 2005; Levy et al., 2005). The KOPS sequence acts as a recognition loading site for FtsK and “determines” the directionality of translocation, which is towards the XerCD-dif site. With that, the directional translocation of DNA substrate is sequence dependent; furthermore, an active translocating FtsK appears to ignore further KOPS sequences and reads through them (Lowe et al., 2008; Lee et al., 2012). Like FtsK, the conserved C-terminal domain of SpoIIIE also harbors three subdomains of α, β, and γ (Fiche et al., 2013; Kaimer & Graumann, 2011), which hexamerize and accommodate the dsDNA substrate in its central channel ring.

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Aker, J., Hesselink, R., Engel, R., Karlova, R., Borst, J. W., Visser, A. J. W. G., de Vries, S. C., 2007. In vivo hexamerization and characterization of the Arabidopsis AAA ATPase CDC48A complex using forster resonance energy transfer-fluorescence lifetime imaging microscopy and fluorescence correlation spectroscopy. Plant Physiology 145, 339–350. Bazinet, C., King, J., 1985. The DNA translocation vertex of dsDNA bacteriophages. Annual Review of Microbiology 39, 109–129. Bigot, S., Saleh, O. A., Lesterlin, C., Pages, C., El, K. M., Dennis, C., Grigoriev, M., Allemand, J. F., Barre, F. X., Cornet, F., 2005. KOPS: DNA motifs that control E. coli chromosome segregation by orienting the FtsK translocase. EMBO Journal 24, 3770–3780. Cardarelli, L., Lam, R., Tuite, A., Baker, L. A., Sadowski, P. D., Radford, D. R., Rubinstein, J. L., Battaile, K. P., Chirgadze, N., Maxwell, K. L., Davidson, A. R., 2010. The crystal structure of bacteriophage HK97 gp6: defining a large family of head-tail connector proteins. Journal of Molecular Biology 395, 754–768. Carrascosa, J. L., Vinuela, E., Garcia, N., Santisteban, A., 1982. Structure of the head-tail connector of bacteriophage phi29. Journal of Molecular Biology 154, 311–324. Caspar, D. L. D., Klug, A., 1962. Physical principles in the construction of regular viruses. Cold Spring Harbor Symposium on Quantitative Biology 27, 1–24. Chen, C., Trottier, M., Guo, P., 1997. New approaches to stoichiometry determination and mechanism investigation on RNA involved in intermediate reactions. Nucleic Acids Symposium Series 36, 190–193. Chen, Y. J., Yu, X., Egelman, E. H., 2002. The hexameric ring structure of the Escherichia coli RuvB branch migration protein. Journal of Molecular Biology 319, 587–591. Chistol, G., Liu, S., Hetherington, C. L., Moffitt, J. R., Grimes, S., Jardine, P. J., Bustamante, C., 2012. High degree of coordination and division of labor among subunits in a homomeric ring ATPase. Cell 151, 1017–1028. Demarre, G., Galli, E., Barre, F. X., 2013. The FtsK family of DNA pumps. Advances in Experimental Medicine and Biology 767, 245–262. Doan, D. N., Dokland, T., 2007. The gpQ portal protein of bacteriophage P2 forms dodecameric connectors in crystals. Journal of Structural Biology 157, 432–436. Dubarry, N., Possoz, C., Barre, F-X. 2010. Multiple regions along the Escherichia coli FtsK protein are implicated in cell division. Molecular Microbiology 78(5), 1088–1100. Fiche, J. B., Cattoni, D. I., Diekmann, N., Langerak, J. M., Clerte, C., Royer, C. A., Margeat, E., Doan, T., Nollmann, M., 2013. Recruitment, assembly, and molecular architecture of the SpoIIIE DNA pump revealed by superresolution microscopy. PLoS Biology. 11, e1001557. Grainge, I., Lesterlin, C., Sherratt, D. 2011. Activation of XerCD- dif recombination by the FtsK DNA translocase. Nucleic Acids Research 39, 5140–5148. Guo, P., Zhang, C., Chen, C., Trottier, M., Garver, K., 1998. Inter-RNA interaction of phage phi29 pRNA to form a hexameric complex for viral DNA transportation. Molecular Cell 2, 149–155. Hendrix, R. W., 1978. Symmetry mismatch and DNA packaging in large bacteriophages. Proceedings of the National Academy of Sciences of the United States of America 75, 4779–4783. Hendrix, R. W., 1998. Bacteriophage DNA packaging: RNA gears in a DNA transport machine (minireview). Cell 94, 147–150. Ibarra, B., Caston, J. R., Llorca, O., Valle, M., Valpuesta, J. M., Carrascosa, J. L., 2000. Topology of the components of the DNA packaging machinery in the phage phi29 prohead. Journal of Molecular Biology 298, 807–815. Iyer, L. M., Leipe, D. D., Koonin, E. V., Aravind, L., 2004. Evolutionary history and higher order classification of AAA plus ATPases. Journal of Structural Biology 146, 11–31. Kaimer, C., Graumann, P. L., 2011. Players between the worlds: multifunctional DNA translocases. Current Opinion in Microbiology 14, 719–725. Kochan, J., Carrascosa, J. L., Murialdo, H., 1984. Bacteriophage lambda preconnectors: Purification and structure. Journal of Molecular Biology 174, 433–447. Lee, J. Y., Finkelstein, I. J., Crozat, E., Sherratt, D. J., Greene, E. C. 2012. Single-molecule imaging of DNA curtains reveals mechanisms of KOPS sequence targeting by the DNA translocase FtsK. Proceedings of the National Academy of Sciences USA 109, 6531–6536. Levy, O., Ptacin, J. L., Pease, P. J., Gore, J., Eisen, M. B., Bustamante, C., Cozzarelli, N. R., 2005. Identification of oligonucleotide sequences that direct the movement of the Escherichia coli FtsK translocase. Proceedings of the National Academy of Sciences of the United States of America 102, 17618–17623. Löwe, J., Ellonen, A., Allen, M. D., Atkinson, C., Sherratt, D. J., Grainge, I. 2008. Molecular mechanism of sequence-directed DNA loading and translocation by FtsK. Molecular Cell 31(4), 498–509.

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Massey, T.H., Mercogliano, C.P., Yates, J., Sherratt, D.J., Lowe, J. 2006. Double-stranded DNA translocation: structure and mechanism of hexameric FtsK. Molecular Cell 23, 457–469. Moll, D., Guo, P., 2007. Grouping of ferritin and gold nanoparticles conjugated to pRNA of the phage phi29 DNA-packaging motor. Journal of Nanoscience and Nanotechnology (JNN) 7, 3257–3267. Morais, M. C., Koti, J. S., Bowman, V. D., Reyes-Aldrete, E., Anderson D, Rossman, M. G., 2008. Defining molecular and domain boundaries in the bacteriophage phi29 DNA packaging motor. Structure 16, 1267–1274. Morais, M. C., Tao, Y., Olsen, N. H., Grimes, S., Jardine, P. J., Anderson, D., Baker, T. S., Rossmann, M. G., 2001. Cryoelectron-microscopy image reconstruction of symmetry mismatches in bacteriophage phi29. Journal of Structural Biology 135, 38–46. Mueller-Cajar, O., Stotz, M., Wendler, P., Hartl, F. U., Bracher, A., Hayer-Hartl, M., 2011. Structure and function of the AAA+ protein CbbX, a red-type Rubisco activase. Nature 479, 194–199. Schwartz, C., De Donatis, G. M., Fang, H., Guo, P., 2013. The ATPase of the phi29 DNA-packaging motor is a member of the hexameric AAA+ superfamily. Virology 443, 20–27. Shu, D., Zhang, H., Jin, J., Guo, P., 2007. Counting of six pRNAs of phi29 DNA-packaging motor with customized single molecule dual-view system. EMBO Journal 26, 527–537. Simpson, A. A., Tao, Y., Leiman, P. G., Badasso, M. O., He, Y., Jardine, P. J., Olson, N. H., Morais, M. C., Grimes, S., Anderson, D. L., Baker, T. S., Rossmann, M. G., 2000. Structure of the bacteriophage phi29 DNA packaging motor. Nature 408, 745–750. Trottier, M., Guo, P., 1997. Approaches to determine stoichiometry of viral assembly components. Journal of Virology 71, 487–494. Valpuesta, J. M., Fujisawa, H., Marco, S., Carazo, J. M., Carrascosa, J., 1992. Three-dimensional structure of T3 connector purified from overexpressing bacteria. Journal of Molecular Biology 224, 103–112. Wang, F., Mei, Z., Qi, Y., Yan, C., Hu, Q., Wang, J., Shi, Y., 2011. Structure and mechanism of the hexameric MecA-ClpC molecular machine. Nature 471, 331–335. Willows, R. D., Hansson, A., Birch, D., Al-Karadaghi, S., Hansson, M., 2004. EM single particle analysis of the ATP-dependent BchI complex of magnesium chelatase: an AAA(+) hexamer. Journal of Structural Biology 146, 227–233. Xiao, F., Zhang, H., Guo, P., 2008. Novel mechanism of hexamer ring assembly in protein/RNA interactions revealed by single molecule imaging. Nucleic Acids Research 36, 6620–6632. Yu, X.C., Tran, A.H., Sun, Q., Margolin, W. 1998. Localization of cell division protein FtsK to the Escherichia coli septum and identification of a potential N-terminal targeting domain. Journal of Bacteriology 180, 1296–1304. Yu, J., Moffitt, J., Hetherington, C. L., Bustamante, C., Oster, G., 2010. Mechanochemistry of a viral DNA packaging motor. Journal of Molecular Biology 400, 186–203. Zhang, F., Lemieux, S., Wu, X., St.-Arnaud, S., McMurray, C. T., Major, F., Anderson, D., 1998. Function of hexameric RNA in packaging of bacteriophage phi29 DNA in vitro. Molecular Cell 2, 141–147. Zhang, H., Endrizzi, J. A., Shu, Y., Haque, F., Sauter, C., Shlyakhtenko, L. S., Lyubchenko, Y., Guo, P., Chi, Y. I., 2013. Crystal structure of 3WJ core revealing divalent ion-promoted thermostability and assembly of the phi29 hexameric motor pRNA. RNA 19, 1226–1237.

4

Structure of Rotation Motors Peixuan Guo and Zhengyi Zhao University of Kentucky

CONTENTS 4.1 Structure of Flagellar Motors ................................................................................................. 59 4.2 Structure of FoF1 ATPase ........................................................................................................ 59 References ........................................................................................................................................ 61

4.1

STRUCTURE OF FLAGELLAR MOTORS

Flagellar motors are composed of two parts: the rotating part, or rotor, and the non-rotating part, or stator (basal body). The rotor is made of four rings (C, coaxial MS, L, and P) and the rod. It is connected to the filament and hook (tubular structure at the base region) and anchored in the cytoplasmic membrane and cell wall (Terashima et al., 2008; Thormann & Paulick, 2010). The stator is a membrane-embedded energy converter composed of MotA/MotB complex with around 12–14 subunits that are organized around two rings in the cytoplasmic membrane. Ion flow through stator (MotAB) is believed to be coupled with force generation, generated by the interaction between rotor and stator components (Terashima et al., 2008; Thormann & Paulick, 2010). Rotation of the motor can be in both directions of clockwise and counterclockwise, and the direction can be switched spontaneously with the complex arrangement of the rotor.

4.2

STRUCTURE OF FOF1 ATPase

FoF1 ATPase, located in the mitochondrial, bacterial, and chloroplast membranes, consists of two rotors, motors termed F1 and Fo. They are located on opposite ends of the stator, a drive shaft known as γ, which connects the motors. Because they are connected, they work as a unit: one motor can cause the other to turn. As both motors share the γ shaft, they compete for control over it. When a steep proton flow moves through the Fo motor, it can force the F1 motor to rotate by turning the γ axle and creating torque, forcing it to act as a generator and thus to produce ATP. This motor is composed of five subunits known as α, β, γ, δ, and ε. At least three α and three β, which are arranged in a hexameric ring around the γ subunit, are required to sustain ATP synthesis. Fo has at least three subunits, polypeptides referred to as A, B, and C. Three ring-shaped C subunits form its rotor portion. F1 and Fo are distinct in structure and function. F1 can be reversibly dissociated from Fo, and shows strong ATP hydrolysis activity that is coupled with inner subunit rotation. Thereby, F1 is termed F1-ATPase. F1 consists of α3β3γ1δ1ε1 subunits in bacterial systems, and its minimum motor complex is the α3β3γ subcomplex (Figure 4.1). The catalytic reaction centers are located at the α-β interface, mainly on the β subunit. Crystal structure revealed that three α and three β subunits form the hexameric stator ring with a large central cavity, which accommodates half of the long coiledcoil structure of the rotary γ subunit (Figure 4.1) (Abrahams et al., 1994). The reported structure of the F1-Fo c complex of yeast ATP synthase showed that the other half of the coiled-coil of γ extends to bind to the rotor part of Fo (the oligomer ring of the c subunits) (Stock et al., 1999). The ε subunit binds to the side surface of the protruding part of the γ and also has close contact with Fo c. The δ subunit is located on the bottom tip of the α3β3 ring. The rotation of γ is the result of cooperative conformational change among the β subunits, exerting the catalysis (Uchihashi et al., 2011). These DOI: 10.1201/9780429203367-5

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FIGURE 4.1 Structures of FoF1 ATP synthase and the α3β3γ subcomplex of F1

catalytic sites can adopt different binding states. The first binds with Mg-AMP-PNP (adenosine 5’-(β, γ-imino)-triphosphate), the second with Mg-ADP, and the third is empty, each referred as to βATP, βADP, and βempty. The two ligand-bound states of β (βATP and βADP) show a closed conformation, swinging the upwardly protruding C-terminal domain toward the central γ subunit (Figure 4.1). These conformational states are termed closed conformation. On the other hand, βempty assumes the opened conformation with the C-terminal domain opening outwardly. Fo is composed of a1b1c8–15 subunits (Figure 4.1). The number of the c subunit differs among species, while the model bacterial Fo has 10 c subunits. Fo generates torque coupled with proton translocation across a membrane down pmf. Compared with F1, Fo is less studied. This is mainly because Fo is a membrane-embedded complex and a highly hydrophobic protein complex, causing difficulties in purification, handling, and activity measurement. The atomic structure of the whole complex of Fo is still not available, although the isolated oligomer ring of the c subunit was crystallized for several species (Stock et al., 1999; Preiss et al., 2013; Vollmar et al., 2009; Pogoryelov et al., 2009; Meier et al., 2005; Watt et al., 2010) (Figure 4.1). While F1 has a pseudo-circular symmetry in the stator ring, Fo has a circular symmetry in the rotor part; the c subunits form the oligomeric ring that rotates against the stator a1b2 complex of Fo, which forms the stator complex with the α3β3 ring in the whole complex of ATP synthase. The proton translocation channel is formed by the a subunit; the c oligomeric ring, i.e., protons, is translocated between the rotor and stator parts. It is thought that upon each proton translocation, the c ring rotates against the a subunit by the angle for a single c subunit in the c ring: 36° steps for the c10 ring. In the whole complex of FoF1 ATP synthase, the c ring binds to the γε complex to form the common rotary shaft of ATP synthase (Stock et al., 1999). The b dimer complex spans the lipid membrane extending toward the headpiece of F1 to bind to the δ subunit that sits on the bottom hollow of the α3β3 ring (Walker & Dickson, 2006). The b2-δ complex forms the peripheral stalk to hold the stator parts of F1 and Fo (the α3β3 ring and the a subunit) stably to transmit the torque efficiently with each other. To achieve the energy interconversion between pmf and free energy of ATP hydrolysis,

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ΔGATP, the rotation directions of Fo are opposite to those of F1. The rotation direction of FoF1 ATP synthase is determined by the magnitude relation of driving forces of Fo and F1, each derived from pmf and ΔGATP. The driving force is defined as the number of driving reactions per rotation divided by 2π. Therefore, when n ⋅ pmf > −3 ⋅ ∆GATP , where n represents the number of c subunits in the c ring of Fo, Fo governs the rotation, reversibly rotating the common rotary shaft (the γε-c ring complex) against F1 that induces the reverse reaction of ATP hydrolysis: ATP synthesis. When n ⋅ pmf < −3 ⋅ ∆GATP , F1 rotates the rotary shaft, enforcing Fo to pump protons to build pmf. Thus, n is physiologically critical to determine the chemical equilibrium of ATP synthesis/hydrolysis. It is thought that n has been tuned under evolutional pressure to meet the physiological requirements (von Ballmoos et al., 2009). Actually, Fo has only eight copies of the c subunit in mammalian mitochondria where respiratory chains produce large pmf (Watt et al., 2010). On the other hand, in some bacteria and chloroplast, of which pmf is supposed to be low, n is large: 13–15 (Preiss et al., 2013; Vollmar et al., 2009; Pogoryelov et al., 2009; Preiss et al., 2014). From a mechanistic point of view, n would be important for smooth energy transmission. In the majority of species studied so far, n is a non-integer multiple of 3, such as 8, 10, and 11. To date, a 9-mer ring has not been found. One rare exception is the 15-mer ring of Fo from Spirulina platensis (Pogoryelov et al., 2009). A rotor ring with threefold symmetry was also found in a bacterial type of vacuole-type H+-ATPase (Toei et al., 2007) that shares remarkably similar structure and working mechanism, although there are some significant differences. Thus, a non-integer multiple of 3 is not an obligation for rotary ATPases. However, the apparent preference of non-threefold symmetry in the proton-conducting unit would represent the intrinsic nature of the rotary coupling mechanism of ATP synthase. One intriguing and reasonable explanation for this is that it would provide a relatively smooth rotary potential surface for the rotor complex, allowing the γε-c ring complex to avoid being trapped in a deep potential minimum where the potential minimums of F1 and Fo overlap.

REFERENCES Abrahams, J. P., Leslie, A. G., Lutter, R., Walker, J. E., 1994. Structure at 2.8 Å resolution of F1-ATPase from bovine heart mitochondria. Nature 370, 621–628. Meier, T., Polzer, P., Diederichs, K., Welte, W., Dimroth, P., 2005. Structure of the rotor ring of F-type Na+ATPase from Ilyobacter tartaricus. Science 308, 659–662. Pogoryelov, D., Yildiz, O., Faraldo-Gomez, J. D., Meier, T., 2009. High-resolution structure of the rotor ring of a proton-dependent ATP synthase. Nature Structural & Molecular Biology 16, 1068–1088. Preiss, L., Klyszejko, A. L., Hicks, D. B., Liu, J., Fackelmayer, O. J., Yildiz, O., Krulwich, T. A., Meier, T., 2013. The c-ring stoichiometry of ATP synthase is adapted to cell physiological requirements of alkaliphilic Bacillus pseudofirmus OF4. Proceedings of the National Academy of Sciences of the United States of America 110, 7874–7879. Preiss, L., Langer, J. D., Hicks, D. B., Liu, J., Yildiz, O., Krulwich, T. A., Meier, T., 2014. The c-ring ion binding site of the ATP synthase from Bacillus pseudofirmus OF4 is adapted to alkaliphilic lifestyle. Molecular Microbiology 92, 973–984. Stock, D., Leslie, A. G., Walker, J. E., 1999. Molecular architecture of the rotary motor in ATP synthase. Science 286, 1700–1705. Terashima, H., Kojima, S., Homma, M., 2008. Flagellar motility in bacteria structure and function of flagellar motor. International Review of Cell and Molecular Biology 270, 39–85. Thormann, KM., Paulick, A., 2010. Tuning the flagellar motor. Microbiology 156(Pt 5), 1275–1283. Toei, M., Gerle, C., Nakano, M., Tani, K., Gyobu, N., Tamakoshi, M., Sone, N., Yoshida, M., Fujiyoshi, Y., Mitsuoka, K., Yokoyama, K., 2007. Dodecamer rotor ring defines H+/ATP ratio for ATP synthesis of prokaryotic V-ATPase from Thermus thermophilus. Proceedings of the National Academy of Sciences of the United States of America 104, 20256–20261. Uchihashi, T., Iino, R., Ando, T., Noji, H., 2011. High-speed atomic force microscopy reveals rotary catalysis of rotorless F-1-ATPase. Science 333, 755–758. Vollmar, M., Schlieper, D., Winn, M., Buchner, C., Groth, G., 2009. Structure of the c14 rotor ring of the proton translocating chloroplast ATP synthase. Journal of Biological Chemistry 284, 18228–18235.

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von Ballmoos, C., Wiedenmann, A., Dimroth, P., 2009. Essentials for ATP synthesis by F1F0 ATP synthases. Annual Review of Biochemistry 78, 649–672. Walker, J. E., Dickson, V. K., 2006. The peripheral stalk of the mitochondrial ATP synthase. Biochimica et Biophysica Acta-Bioenergetics 1757, 286–296. Watt, I. N., Montgomery, M. G., Runswick, M. J., Leslie, A. G. W., Walker, J. E., 2010. Bioenergetic cost of making an adenosine triphosphate molecule in animal mitochondria. Proceedings of the National Academy of Sciences of the United States of America 107, 16823–16827.

5

Structure of Linear Motors Peixuan Guo and Zhengyi Zhao University of Kentucky

CONTENTS 5.1 Structure of Myosins............................................................................................................... 63 References ........................................................................................................................................64

5.1

STRUCTURE OF MYOSINS

Most linear motors possess head, neck, and tail domains, with the head allowing and directing movement, the tail identifying its cargo, and the neck connecting the head and tail. Unlike the other two types of motors, they contain all the elements capable of converting chemical energy into mechanical work. Myosin, kinesin, and dynein power many different essential functions. The structure of the activity domains of linear motors (Figure 5.1) is highly conserved among various classes of myosins (Geeves & Holmes, 1999; Vale & Milligan, 2000). Therefore, the conformational pathway of the myosin ATPase cycle is hypothesized to be similar in all myosin motors, while variation in the kinetic and equilibrium constants of the conformational changes allows for myosins to be fine-tuned for performing specific cellular functions (De la Cruz & Ostap, 2004). The motor domain contains ATP-binding (nucleotide binding pocket—NBP) and actin-binding (actin binding cleft-cleft) motifs (Sweeney & Houdusse, 2010), which are coupled with the reciprocal movement of the lever-arm region during the recovery and power stroke states of the ATPase cycle (Trivedi, 2014) (Figure 5.1). However, the mechanism of allosteric communication between different subdomains of the motor remains a crucial question in the field today. Any perturbation to these communication pathways has been hypothesized to lead to pathophysiological conditions. While the allosteric communication between the nucleotide- and actin-binding regions has been extensively studied (Málnási-Csizmadia et al., 2005; Kintses et al., 2007; Conibear et al., 2003), there remain outstanding questions regarding the coupling of actin binding, product release, and the position of the lever arm (Malnasi-Csizmadia & Kovacs, 2010). A long-standing question in the actomyosin field is the precise timing of force generation and its relationship to the kinetics of lever-arm swing. The mechanical work produced by myosin includes muscle contraction, movement of cargo or organelles on actin filaments, membrane tension generation, endocytosis, and exocytosis. They can also participate in signal transduction and transcription. Kinesin and dynein carry out much of the long-distance transport in neurons as well as a diversity of other functions such as the bending of cilia and flagella. Kinesin and myosin share a similar structural fold and are members of the P-loop NTPases that contain a conserved nucleotide-binding region. The overall organization of their motor domains is similar with the conserved nucleotide-binding region, which serves as the motor core that communicates with the microtubule- or actin-binding region as well as the forcegenerating elements. Dynein is a more complex ring-shaped motor that has six ATP-binding regions and is a member of the AAA+ family. Only four of the six nucleotide-binding sites bind nucleotide, while a single site controls the movement. Myosin V (MV) is an unconventional, dimeric, and highly processive myosin involved in transporting vesicles and cargoes along actin filaments in cells (Mehta et al., 1999). It can walk along actin filaments using a hand-over-hand mechanism, taking 36-nm steps to transport an associated cargo (Yildiz et al., 2003). The overall structure of MV consists of a motor domain that has DOI: 10.1201/9780429203367-6

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FIGURE 5.1 Crystal structure showing the different subdomains of myosin along with the actin, nucleotide binding, and lever arm regions (PDB code 1W7J).

actin- and nucleotide-binding regions, followed by a neck and tail region that allows dimerization and cargo binding. The neck region (lever arm) of MV has 6 IQ-domains that can individually bind calmodulin (CaM) and mechanically stiffen the lever arm to stabilize the position of the motor domain for coordinated stepping. MV has served as an outstanding model to study actin-induced structural changes as its affinity for actin is much higher in the weak binding states compared to conventional myosin II (Yengo et al., 2002). This feature allows MV to stay attached to actin at lower actin concentrations, and structural differences between the weak and strong actin-binding states can be elucidated with spectroscopic, kinetic, and structural studies (Jacobs et al., 2011; Veigel et al., 2005; Coureux et al., 2004).

REFERENCES Conibear, P. B., Bagshaw, C. R., Fajer, P. G., Kovacs, M., Malnasi-Csizmadia, A., 2003. Myosin cleft movement and its coupling to actomyosin dissociation. Nat Struct Biol 10, 831–835. Coureux, P. D., Sweeney, H. L., Houdusse, A., 2004. Three myosin V structures delineate essential features of chemo-mechanical transduction. EMBO J 23, 4527–4537. De la Cruz, E. M., Ostap, E. M., 2004. Relating biochemistry and function in the myosin superfamily. Curr Opin Cell Biol 16, 61–67. Geeves, M. A., Holmes, K. C., 1999. Structural mechanism of muscle contraction. Annu Rev Biochem 68, 687–728. Guo, P., Noji, H., Yengo, C. M., Zhao, Z., Grainge, I., 2016. Biological nanomotors with revolution, linear, or rotation motion mechanism. Microbiol Mol Biol Rev 80, 161–186. Jacobs, D. J., Trivedi, D., David, C., Yengo, C. M., 2011. Kinetics and thermodynamics of the rate-limiting conformational change in the actomyosin V mechanochemical cycle. J Mol Biol 407, 716–730. Kintses, B., Gyimesi, M., Pearson, D. S., Geeves, M. A., Zeng, W., Bagshaw, C. R., Malnasi-Csizmadia, A., 2007. Reversible movement of switch 1 loop of myosin determines actin interaction. EMBO J 26, 265–274. Málnási-Csizmadia, A., Dickens, JL., Zeng, W., Bagshaw, CR., 2005. Switch movements and the myosin crossbridge stroke. J Muscle Res Cell Motil 26, 31–37. Malnasi-Csizmadia, A., Kovacs, M., 2010. Emerging complex pathways of the actomyosin powerstroke. Trends Biochem Sci. 35, 684–690.

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Mehta, A. D., Rock, R. S., Rief, M., Spudich, J. A., Mooseker, M. S., Cheney, R. E., 1999. Myosin-V is a processive actin-based motor. Nature 400, 590–593. Sweeney, H. L., Houdusse, A., 2010. Structural and functional insights into the myosin motor mechanism. Annu Rev Biophys 39, 539–557. Trivedi, D. V., 2014. Allosteric communication and force generation in myosin motors. Doctoral dissertation. Pennsylvania State University, University Park, PA. Vale, R. D., Milligan, R. A., 2000. The way things move: looking under the hood of molecular motor proteins. Science. 288, 88–95. Veigel, C., Schmitz, S., Wang, F., Sellers, J. R., 2005. Load-dependent kinetics of myosin-V can explain its high processivity. Nat Cell Biol 7, 861–869. Yengo, C. M., De la Cruz, E. M., Safer, D., Ostap, E. M., Sweeney, H. L., 2002. Kinetic characterization of the weak binding states of myosin V. Biochemistry 41, 8508–8517. Yildiz, A., Forkey, J. N., McKinney, S. A., Ha, T., Goldman, Y. E., Selvin, P. R., 2003. Myosin V walks handover-hand: single fluorophore imaging with 1.5-nm localization. Science 300, 2061–2065.

6

Mechanical Properties of Molecular Motors and the Relevance to Their Biological Function Yuchuan Zheng and Jingyuan Li Zhejiang University

CONTENTS 6.1 Kinesin .................................................................................................................................... 68 6.2 Myosin .................................................................................................................................... 70 6.3 F0F1-ATPase ............................................................................................................................ 71 6.4 Φ29 DNA Packaging Motor ................................................................................................... 73 References ........................................................................................................................................ 76 Molecular motors are molecular devices possessed by cells that can convert chemical energy derived from ATP hydrolysis into mechanical work [1]. Molecular motors are involved in a wide array of biological processes, including cell movement, intracellular transport, ion gradient generation, and protein folding and unfolding [2–6]. There are two important characteristics that highlight the performance of the molecular motor, i.e., energy efficiencies and power-to-weight ratios. Energy efficiency refers to the ratio of the mechanical work output to the energy consumed in one cycle of the motor; power-to-weight ratio refers to the ratio of the output power of the motor to its own mass. From both perspectives, the molecular motors outperform most man-made motors. The energy efficiency of molecular motor is usually much higher than those of the most man-made motors. For example, the F0F1-ATPase features that the energy efficiencies are up to almost 100% [7]. On the other hand, the energy efficiencies of common man-made motors such as gasoline engine and diesel engine are usually less than 36% and 44% [8]. Similarly, a variety of biological molecular motors exhibit exceptional power-to-weight ratio as compared to man-made motor. For example, the powerto-weight ratio of kinesin can be up to 23.4 kW/kg [9], whereas the ratio of car engine (e.g., Mazda 13B-MSP Renesis 1.3 L Wankel) is only 1.5 kW/kg [10] and the ratio of jet engine (e.g., GE90-115B Brayton turbofan) is 10.0 kW/kg [11,12]. It is interesting to note that molecular motor works in ambient temperature and pressure, whereas the rocket engine (e.g., Top Fuel Dragster Engine, 36.46 kW/ kg [13]) works in extremely high temperature. Such remarkable energy efficiencies of molecular motors imply that the force can be effectively transmitted throughout the structure of molecular motor. In addition, the superior power-to-weight ratio infers that molecular motors need to withhold exceptional workloads and sustain their structure integrity. Taken together, the mechanical properties of molecular motors are directly related to their performance and have significant biological relevance. It should be noted that most molecular motors consist of multiple components. And there are a large variety of assembly architectures of molecular motors. Moreover, the operation modes of molecular motor are also miscellaneous. The motors can thus be largely divided into three categories, i.e., translational, rotational, and DOI: 10.1201/9780429203367-7

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revolutionary motors. Hence, the resulting distribution of mechanical strain within motor’s structure should be highly diversified. Comprehensive understanding of mechanical properties of molecular motors as well as the relationship with their performance demands systematic interpretation of the mechanical properties of these components, which should shed light to the structural basis of the effective force transmission and the mechanical resilience throughout the structure of molecular motor. With the help of state-of-art techniques including atomic force microscopes [14,15], optical traps [16,17], glass needles [18], there are growing number of studies about the mechanical properties of molecular motors as well as their components. In this chapter, we review the studies about the mechanical properties of a series of representative molecular motors of each category, including kinesin, myosin (translational motor), F0F1-ATPase (rotational motor), and the Φ29 DNA packing motor (revolutionary motor).

6.1

KINESIN

Kinesin is a molecular motor that transports intracellular cargoes including membrane organelles, mRNAs, and protein complexes along the microtubules (MTs) by exploiting the energy derived from ATP hydrolysis. Kinesin travels along the MTs with step-like motions by means of two catalytic motor domains (i.e., heads) binding and unbinding alternatively on the MT [2,4,19–21]. Both heads are joined together by 4-nm-long segments called neck linkers to the N terminus of an extended stalk, which consists of three stiff coiled coils (CC) (i.e., neck CC, CC 1 and 2) connected by two flexible regions called hinge (i.e., hinge 1 and 2). The C-terminal end of the stalk is a globular domain called tail, which is responsible for binding to various cargoes [22] (Figure 6.1).

FIGURE 6.1

A schematic diagram of the kinesin bound to a microtubule with associated light chains [15].

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Kinesin generates about 5-pN force in its walking step along the MT, and the stall force of kinesin is found to range from 5 to 8 pN [23–26]. Hence, there should be considerable strain distributed throughout the structure of kinesin during operation. The mechanical properties of kinesin are directly related to its function. There are a variety of biophysical studies to estimate the mechanical properties of the components of kinesin. As indicated by these studies, the mechanical stabilities of various kinesin components are highly diversified. For example, the CC regions are very stiff: the stretching stiffness of the full 70-nm CC is up to 85 pN/nm, as calculated in H. Grubmuller’s computational study [27]. As estimated by atomic force microscopy, CC can remain folded under the transverse force up to 11 pN [28]. On the other hand, the hinges are much more flexible. S. M. Block and coworker found that the torsional stiffness of hinges is much weaker than that of the CC domains, with Young’s modulus of 0.02 and 0.6 GPa [29], respectively. Moreover, the mechanical stability of kinesin’s component appears to be sensitive to ATP binding. For example, neck linker is quite flexible in the absence of ATP serving as a connector between the heads and the neck CC. However, neck linker becomes rigid upon ATP binding and “docks” to the head domain forcing the connection point of head to stalk forward by ∼2.7 nm [22,26]. Such conformational change of neck linker is thus essential for kinesin’s moving on the MT, which will be discussed below. As indicated by previous studies, each ATP hydrolysis causes kinesin an 8-nm step-like motion along the MT [21]. Initially, only one head (denoted as leading head) binds on the MT, and the other head (denoted as moving head) diffuses behind [30]. Binding of ATP triggers a conformational change in leading head, resulting in the folding of a β-sheet formed by the N-terminal of leading head and neck linker [31]. Such conformational change can engender a forward conformational bias to overcome mechanical loads and drives the moving head forward motion. The moving head is then guided to the next binding site through its electrostatic interaction with MT [32]. After the moving head’s binding on the MT, the strain between two bound heads becomes up to 12 ~ 15 pN [33]. By means of such considerable strain, kinesin effectively coordinates the walking pace of two heads. The leading head detaches from the MT after sensing such strain on neck linker, and becomes the moving head in the next stepping cycle. As discussed above, mechanical stabilities of various components of kinesin are highly diversified. Moreover, the heterogeneous nature of the mechanical stability throughout kinesin’s structure is highly related to the responsibilities of various components during operation and is thus essential for kinesin’s function. For example, the stiff CC facilitates the force transmission from the motor heads to the cargo-bounded tail region, contributing to the high efficiency of kinesin. In addition, the flexible hinges can promptly relieve strains that develop during the motion of kinesin [29]. Hence, kinesin can relieve the twist accompanied by cargo transport while maintaining both directionality and processivity of stepping motion [29]. The neck linker with ATP-sensitive mechanical property plays a key role in the force generation and the coordination of the behaviors of two heads. On the one hand, as a key component of kinesin’s power generation [31], the rigid structure of neck linker upon ATP binding enables itself to effectively generate the force that drives kinesin forward. In addition, neck linker is also essential for kinesin to coordinate its two identical motor domains. The strain on neck linker regulates MT dissociation of the leading head as well as the two heads’ enzymatic cycle [33]. ADP release from the moving head is repressed until it is propelled to a forward position triggered by neck linker docking in the leading head [34,35]. Moreover, the subsequent ATP binding to the MT-bound front moving head is inhibited until the leading head dissociates from the MT [36,37]. Through the effective strain conduction on the neck linker, kinesin coordinates the activities of its two heads and keeps them out of phase. Therefore, kinesin can realize both tight chemomechanical coupling (i.e., each ATP hydrolysis causing a step-like motion) and high processivity [33].

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MYOSIN

Myosin II is a representative member of myosin family and is essential for muscle contraction. With its catalytic motor domains binding on the actin filaments, myosin II hydrolyzes ATP to generate the force that drives the forward sliding movement of actin filaments during muscle contraction [38–41]. Myosin II can be divided by proteolytic cleavage into three fragments: S1, S2, and LMM (i.e., light meromyosin) [42–46]. S1 fragment consists of two motor heads and two pairs of light chains (i.e., ELC and ERC), and is the domain responsible for ATP hydrolysis and force generation. S2 fragment is a coiled coil that connects the S1 fragment to the LMM-CC tail. The LMM-CC tail further joins to the myosin filament, an assembly of multiple myosin II molecules that slide relative to the actin filaments during muscle contraction [39,41] (Figure 6.2). Myosin II shares similar function with kinesin that discussed above. Myosin II is also a translational motor and moves along the filaments with two catalytic motor heads. Moreover, the motor domains of kinesin and myosin II show considerable structural homology [47–49]. Their motor domains are formed by similar core structural elements and organized in an analogous way. In addition, both motors share a mechanism of force generation to drive the movement of motor heads along the tracks, i.e., generating force through a conformational change powered by ATP hydrolysis in the catalytic motor heads [26,50]. Despite the similarities in their structural arrangement, there are substantial differences that exist between myosin II and kinesin. For example, myosin II and kinesin hydrolyze ATP in different stages of their mechanical cycles, i.e., after the detachment of myosin II from actin vs. when kinesin is still bound to MTs (50). Besides, the force-generating conformational changes in two motors are triggered in different events in motors’ mechanical cycles. In myosin II, the conformational change is caused by the binding of the motor head to the actin filament. However, the conformational change in kinesin results from the binding of ATP to the bound motor head [26,50]. Moreover, unlike kinesin that walks progressively along the MT, myosin II is a non-progressive motor that detaches from the actin filaments after working strokes [51]. Thus, in order to power the muscle contraction whose duration is much longer than single myosin’s working stroke, the cooperation of various myosin II molecules is highly warranted. Hence, multiple myosin II molecules join together with their tail regions and form the myosin filament (Figure 6.3) [38,41,44]. Through this assembly, these myosin molecules mutually interact with one actin filament. Such arrangement allows occasional single-molecule detachment during muscle contraction and facilitates myosin II to generate force collectively and drive the actin filaments forward [51].

FIGURE 6.2 Schematic representation of the myosin II protein and its subfragments. (Adapted from http:// www.mrothery.co.uk/images/Imag108.gif.) [18].

Mechanical Properties of Molecular Motors

FIGURE 6.3

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Schematic representation of myosin (thick) filaments and actin (thin) filaments [18].

As mentioned above, the muscle contraction is the consequence of sliding movements of actin filaments as driven by the collective behaviors of a great amount of myosin II molecules’ working strokes. The mechanical properties of individual myosin II molecule are directly related to the overall behavior of muscle contraction. The mechanical properties of myosin II have thus been intensively studied. Many studies have measured the stiffness of myosin II molecule based on the assumption that myosin II molecule possesses linear elasticity. Interestingly, stiffness measured in these studies is highly diversified, ranging from 0.17 to 3 pN/nm [52–57]. Such variation in the measured stiffness is largely due to the fact that the elasticity of myosin II under positive and negative strain is actually nonlinear. By using a combination of optical tweezer and single fluorescence imaging technique, H. Higuchi and coworkers found that the stiffness of negatively strained myosin II is substantially smaller than that of the positive strain side (2.9 pN/nm) [53]. Additional computational study demonstrates that such nonlinear elasticity of myosin II is mainly due to its anisotropic S2 fragment. The S2-CC is much stiffer in the axial direction than in the lateral direction, with a stretching stiffness of 60–80 pN/nm and a bending stiffness of ~0.01 pN/nm, respectively [58]. Hence, much lower stiffness of myosin II under negative strain may be related to a bending or buckling of the S2 portion with very small bending stiffness. Nonlinear stiffness of myosin II is essential for the cooperation of multiple myosin II molecules and the overall performance of muscle contraction. During muscle contraction, especially when the actin filament moves at a fast shortening speed, some after-working myosin II molecules remain attached to the actin and are subsequently loaded with the negative strain as the actin is pulled by the other working myosin II [51]. The negatively strained myosin II molecules then interfere with the working of neighboring molecules and engender a drag force that hinders the movement of actin. In such case, the negatively strained myosin II with much lower stiffness is able to reduce such drag force, resulting in the upper limits of the shortening speed of actin and the efficiency of muscle contraction. Besides, the higher stiffness of the positively strained myosin II facilitates the generation of a high working force with a relatively small amount of strain, which also helps in the muscle contraction [51,58].

6.3

F0F1-ATPase

F0F1-ATPase is an essential molecular motor whose function is to synthesize ATP using the energy derived from transmembrane proton motive gradient [5,59–61]. At high ATP concentrations, it can also reverse itself and hydrolyze ATP, serving as a proton pump (Figure 6.4). Different from translational motors like kinesin and myosin II, F0F1-ATPase is the motor that operates through a rotary motion. It consists of two oligomeric portions: the membrane-embedded ion-translocating F0 portion and the catalytic soluble F1 portion [5,59,60,62,63]. The structure of

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FIGURE 6.4 Structure of F0F1-ATPase [16].

rotational F0F1-ATPase is arranged as a “rotor” and “stator” assembly. The stator portion consists of a catalytic α3β3 hexamer, together with subunits a, b, and δ. The α3β3 hexamer is composed of three αβ-subunits arranged as a symmetric ring. Each αβ-subunit has a catalytic site at its αβ-domain interface [62–64]. The rotor portion consists of a ring structure (denoted as c-ring) that composed of 10–15 c-subunits, along with subunits γ and ε [65,66]. The γ-subunit is a connecting component locating in-between the α3β3 hexamer and the c-ring. The upper part of γ-subunit penetrates inside the α3β3 hexamer ring (denoted as α3β3-penetrating portion), and the lower part of γ-subunit is a protruded globular portion that connects to the c-ring and the ε-subunit. In the synthesis mode, the rotor portion rotates relatively to the stator as powered by translocating protons through the interface of the a-subunit and c-ring [5,67–69]. During the rotation motion, the α3β3-penetrating portion of the γ-subunit interacts with the catalytic sites in the α3β3 hexamer and synthesizes ATP from ADP and phosphate. Through this rotary mechanism, F0F1-ATPase can effectively couple the ATP synthesis with proton translocation driven by the transmembrane electrochemical potential gradient. The operation of F0F1-ATPase is also very efficient. It may achieve almost 100% energy efficiency while working steadily at a rotational rate of ~1 kHz [7,70]. In an early study, G. Oster and coworker found that this remarkable performance is related to the effective elastic power transmission between F0 and F1 portions [7]. The energy gained from translocating proton is first converted into torsional strain in the structures of F0F1-ATPase, and the strain is then released to synthesize ATP from ADP and phosphate. Hence, F0F1-ATPase exploits the torsional strain distributed along its structure to couple the proton translocation and ATP synthesis. The torsional elastic properties of F0F1-ATPase should be important to its performance and biological function. There are a variety of biophysical studies about the torsional property of the components of F0F1-ATPase. As shown in these studies, the torsional properties of its components are very heterogeneous. The overall torsional stiffness of the F0F1-ATPase is 68 pN·nm [71]. And the stator is much stiffer, with a torsional stiffness of >500 pN·nm [72]. Such stiff stator portion acts as the scaffold for the ATPase. As for the rotor portion, its torsional stiffness is quite heterogeneous. For example, the α3β3-penetrating portion of γ-subunit is relatively stiff, with a torsional stiffness of 520–750

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pN·nm [63,71]. And the globular portion of the γ-subunit is much complaint, with an elastic module of 120 pN·nm [73]. The interface of the F1 and F0 portion, i.e., the contact region of the c-ring and subunits γ and ϵ, has been identified to be even more complaint: with a torsional stiffness of about 20 pN·nm [72]. The heterogeneous nature of the rotor’s torsional stiffness is essential for the operation of F0F1ATPase. For example, the complaint part of the rotor is able to compensate the rotational symmetry mismatch between F0 and F1. Both F0 and F1 are steppers; the rotational symmetries of F0 and F1 may or may not match with C3 symmetry in F1 and C10–15 symmetry in F0 (depending on organism) [62,74]. The complaint portion of rotor (i.e., globular portion of γ-subunit and the contacting region of F0 and F1) can compensate this symmetry mismatch by temporarily storing the energy gained from single-proton translocation until the rotor has performed sufficient work to drive ATP synthesis [75–77]. This transient energy storage in the complaint part of the rotor smooths the operation of the F0F1-ATPase and contributes to its remarkable energy efficiency. On the other hand, the stiff α3β3-penetrating portion can effectively conduct the strain and interact with the catalytic αβ-subunits. Such structural stiffness facilitates the tight coupling of the two parts of motor and guides them onto a unique rotational pathway [70]. Hence, the motor can work steadily at a high rotational speed. In conclusion, the heterogeneous nature of rotor’s torsional stiffness effectively couples various parts of F0F1-ATPase, contributing to its remarkable performance.

6.4

Φ29 DNA PACKAGING MOTOR

Φ29 DNA packaging motor is a molecular motor exploited by bacteriophage Φ29 to pack DNA into its capsid [78]. Powered by the energy of ATP hydrolysis, Φ29 motor can transport viral DNA against the enormous internal-to-external pressure difference of beyond 50 atm [79,80]. It is thus regarded as one of the most powerful molecular devices in viral, bacterial, and eukaryotic systems alike [80,81]. Assembled at the base of the crowning capsid, Φ29 motor is composed of a head-tail connector, a prohead RNA (pRNA) ring, and an adenosine triphosphatase (ATPase) gp16 ring [82–85]. These components are arranged along the portal axis of capsid. Locating at the portal of capsid, the connector is a dodecameric protein complex with truncated cone shape [86] (Figure 6.5). The connector serves as a channel through which viral DNA can be transported into the capsid. It can be divided into three major regions: upper, middle, and bottom regions. The upper and middle regions are embedded in the capsid, and the bottom region protrudes toward the outside of capsid and connects to the motor’s pRNA component [87]. The pRNA acts as a linker element that joins the connector and ATPase of Φ29 motor. The core structure of pRNA is composed of three major helices organized into a three-way junction (i.e.,

FIGURE 6.5 The structure of the connector: the upper region (blue), middle region (brown), and bottom region (green). The hinge region (red) joins the middle region and bottom region [33].

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FIGURE 6.6 respectively.

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The scheme of the 3WJ-pRNA. RNA strands A, B, and C are colored in red, green, and blue,

3WJ-pRNA) (Figure 6.6) [88–90]. The helix 1 (H1) and helix (H3) of 3WJ-pRNA are stacked coaxially and parallel to the portal axis of motor, while the helix 2 (H2) orients toward the transverse direction. With H1 and H3, respectively, attached to the ATPase- and connector-binding region of pRNA, 3WJ-pRNA effectively links the connector and ATPase domains of Φ29 motor. And H2 is attached to the regions involved in pRNA oligomerization [79,84,88,91,92]. Capable of generating ~20 pN force, Φ29 motor can drive the double-stranded DNA into exceptionally high packing densities [80,81]. During this progress, the motor works against considerable internal-to-external pressure difference induced by DNA confinement [80,93]. Components of Φ29 motor thus bear enormous strain along the portal axis of capsid. Therefore, mechanical properties of the motor’s components are directly related to the performance of motor. There are several biophysical studies to estimate the mechanical properties of the components of Φ29 motor. The motor’s components exhibit diversified mechanical stabilities. For example, the mechanical stability of the connector is quite heterogeneous. The middle region of connector is very rigid and considered as one of the stiffest protein materials [94–96]. As shown in computational study of H. Grubmuller and coworker, Young’s moduli of the middle region is 3.4 ± 0.6 GPa [87], comparable to the protein materials like collagen fibrils (0.2–11.5GPa) [97], single-brin silkworm silk (5–17 GPa) [98], and dragline spider silk (11–13 GPa) [99]. As for the upper region of connector, its top part that is exposed to the capsid interior is very stiff, with similar mechanical property as the middle region. However, such rigid part is embedded within the flexible parts locating at the base of upper region [87]. This arrangement of the stiff and flexible parts of the upper region of connector is similar to those of some composite materials [100–102]. And the bottom region of connector is composed of flexible moieties and is the most complaint region of the connector, with Young’s moduli of less than 0.36 ± 0.06 GPa [87]. The mechanical property of 3WJ-pRNA is directional dependent. As indicated by our molecular dynamics simulation, 3WJ-pRNA exhibits exceptional mechanical stability along the coaxial axis of H1 and H3 (parallel to the portal axis of capsid). When stretched along the portal axis, 3WJ-pRNA can maintain its native structure under the large applied force of 1990 ± 126 pN [103]. This mechanical stability is extraordinary compared to other nucleic acid structures like DNA double helices, RNA hairpins, and pseudoknots that feature a rupture force of a few tens of piconewtons [104–112]. Moreover, this robust mechanical stability is highly directional dependent. 3WJ-pRNA shows little resistance to the applied force and unfolds easily when it is stretched along the transverse direction [103]. Diversified mechanical stability of Φ29 motor’s components is cooperative to their responsibility and crucial for the operation of Φ29 motor. As for the connector, both the middle region and the top part of upper region are exposed to the inner surface of capsid and thus bear considerable pressure during DNA packing. Stiffness of these parts is essential for maintaining the structural integrity of

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the DNA-transporting channel in connector and is crucial for the connector to prevent DNA leakage when the packing proceeds [87]. On the other hand, the flexible parts in the upper region act as buffer between the rigid parts of connector and the capsid, absorbing local force and distributing external force evenly over the interior surface of capsid [87]. It should be noted that complaint bottom region facing the capsid exterior does not need to withstand mechanical stress and pressure difference. Moreover, its high flexibility might facilitate the binding of the other motor components during the packaging process. The asymmetry mechanical stability of pRNA also plays an important role during the operation of Φ29 motor. During the DNA packing progress, the pressure induced by DNA confinement can reach beyond 50 atm along the portal axis of capsid [80,93]. As the linker element between the connector and ATPase domains, the pRNA has to bear considerable mechanical force exerted along this coordination [79,88,91,92,113]. Extraordinary stability along the portal axis is required for pRNA to withstand enormous strain and is crucial for maintaining the structural integrity of Φ29 motor [113]. On the other hand, flexibility in the transverse direction can facilitate the assembly of pRNA and its binding to the capsid prohead [113]. Taken together, heterogeneous nature of Φ29 motor’s mechanical stability is entwined with its biological function: stiffness distributed in the structure of Φ29 motor is necessary to its working in such extreme condition, while flexibility in the motor’s other components also contributes to the motor’s assembly. As discussed above, molecular motors undertake various motions in the cells as driven by chemical energy. For example, F0F1-ATPase operates with its rotor portion that rotates relative to the stator portion [5,59–61]. And kinesin moves translationally with step-like motions performed by two catalytic motor domains (i.e., heads) binding and unbinding alternatively on the MT [2,4,19–21]. Components of these molecular motors exhibit a wide spectrum of mechanical properties. And these highly diversified mechanical properties are directly associated with the motion of motor and contribute to these motors’ remarkable performance. On the one hand, diversified mechanical properties of motors’ components cooperate to the motors’ motion and help these molecular motors to achieve high energy efficiency. For example, the rotor portion of F0F1-ATPase can temporarily store the chemical energy into elastic strains and compensate the rotational symmetry mismatch between F0 and F1 [75–77]. It can thus smooth the ATPase’s operation and result in the motor’s extraordinary energy efficiency. In the case of kinesin, the ATP-sensitive mechanical property of its neck linker can coordinate the walking pace and enzymatic cycle of two motor heads [33], contributing to kinesin’s high processivity and tight chemomechanical coupling. As for myosin II, the anisotropic S2 fragment can buffer the interference of the negatively strained myosin II molecules and reduce the energy loss during muscle contraction [51]. On the other hand, mechanical rigidity within motor’s components sustains motors’ working under large loads and contributes to these motors’ high power-to-weight ratios. For example, the stiff stator in F0F1-ATPase acts as the scaffold that enables the ATPase to work steadily at high rotational speeds [70,72]. And enhanced mechanical stability of Φ29 DNA packaging motor’s 3WJ-pRNA along its portal axis is crucial for the motor to withstand the enormous strain during the DNA packing [103]. Meanwhile, the heterogeneous distribution of mechanical stability in Φ29 DNA packaging motor’s connector portion effectively sustains the structure of DNA transporting channel [87], and is essential for preventing DNA leakage despite the enormous pressure induced by the DNA confinement. Taken together, mechanical properties of molecular motor’s components are highly diversified as discussed above. The heterogeneous mechanical properties of motors’ various components facilitate the operation of motors and contribute to their remarkable performance. Further studies to elucidate the molecular motor’s mechanical properties will help in the understanding of these extraordinary molecule machines and the life processes they participate. Moreover, researches about the structural basis of the mechanical stabilities of molecular motors may shed light on the evolutionary design process of these molecular machines and inspire further development of artificial machines with better performance.

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7

Molecular Mechanism of AAA-ATPase Motor in the 26S Proteasome Shuwen Zhang Peking University

Youdong Mao Peking University

CONTENTS 7.1 7.2 7.3 7.4 7.5

Introduction ............................................................................................................................ 81 AAA+ ATPases in Ubiquitin-Proteasome System ................................................................. 82 Conformational Changes of AAA ATPases in the 26S Proteasome...................................... 85 Substrate Interactions Coupled with ATP Hydrolysis ............................................................ 86 Three Modes of Coordinate ATP Hydrolysis Regulate Intermediate Functional Steps ........ 87 7.5.1 Mode 1 Regulates Ubiquitin Recognition, Initial Substrate Engagement, and Deubiquitylation ......................................................................................................... 88 7.5.2 Mode 2 Regulates CP Gating, Ubiquitin Release, and Initiation of Substrate Translocation............................................................................................... 88 7.5.3 Mode 3 Regulates Processive Substrate Unfolding, Translocation, and Degradation ........ 89 7.6 Evidence for a Sequential Hand-over-Hand Model ...............................................................90 7.7 Concluding Remarks ..............................................................................................................90 Funding ............................................................................................................................................ 91 Acknowledgments............................................................................................................................ 91 Conflicts of Interest .......................................................................................................................... 91 References ........................................................................................................................................ 91

7.1 INTRODUCTION Protein degradation plays a fundamentally pivotal role in the maintenance of cellular homeostasis and the regulation of nearly all major cellular processes, such as cell cycle, gene expression, signal transduction, immune response, apoptosis, and carcinogenesis (Ciechanover, 2005). Not only misfolded, damaged, unneeded, and aggregation-prone proteins, but also regulatory proteins are targeted in the pathways of intracellular proteolysis, dysfunction of which is associated with many human diseases (Ciechanover and Kwon, 2015; Hnia et al., 2019; Mayer, 2000; Meyer-Schwesinger, 2019). In cells, well-folded proteins that are destined for breakdown must be tagged via ubiquitylation pathways to become distinguishable from those normal cellular constituents by the proteasome. The globular domains of these target proteins have to be unfolded and transported to the proteolytically active sites before they can be broken down into short polypeptides. This sophisticated task is thought to be carried out by the ring-like motor modules assembled from adenosine triphosphatase (ATPase) in AAA+ (ATPases associated with a variety of cellular activities) superfamily (Smith et al., 2006; Vale, 2000). AAA+ ATPases are assembled with proteolytic subunits or DOI: 10.1201/9780429203367-8

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FIGURE 7.1 Domain organizations of AAA+ proteases in different families. Length of the bar is not linearly proportional to the real length of the corresponding sequence. Each protein contains one or two AAA+ modules, each consisting of a large and small subdomain, and additional family-specific domains, which are not specifically depicted here. Protease modules reside in separate protein subunits except for FtsH and Lon.

domains into an ATP-fueled complex machinery that couples cyclic ATP hydrolysis with consecutive proteolysis reactions (Arrigo et al., 1988; Dong et al., 2019; Etlinger and Goldberg, 1977). The proteolytic subunits typically assemble into a cylinder-shaped proteolytic chamber, such as proteasome core particle (CP) (Arrigo et al., 1988; Dong et al., 2019; Groll et al., 1997; Lowe et al., 1995), HslV (Bochtler et al., 1997), and ClpP (Wang et al., 1997). In the proteasome, the AAA-ATPase motor is assembled from six distinct of RPT subunits into a heterohexameric ring with a central axial pore and guards the entry port of the CP chamber (Dong et al., 2019). The AAA-ATPase ring is associated with several ubiquitin receptors and deubiquitylating enzyme (DUB) subunits and is further capped with a lid subcomplex to form a regulatory particle (RP) in the 26S proteasome (Arrigo et al., 1988; Dong et al., 2019; Zhang et al., 2022). Powered by the AAA+ ATPase engines, the protease machines can recruit and unfold substrates carrying specific degradation signals and translocate them into the proteolytic chamber for breakdown. The AAA+ superfamily members include several clades (Erzberger and Berger, 2006; Hanson and Whiteheart, 2005; Iyer et al., 2004; Lupas and Martin, 2002; Ogura and Wilkinson, 2001). There are mainly two clades of AAA+ ATPases related to protease complexes. One is the classic AAA clade, including proteasomal family, FtsH family, Cdc48 family, and ClpA/B/C-Domain 1 (D1) family. The other is the HCLR clade, which features a pre-sensor 1 insertion, including HslU/ClpX family, ClpA/B/C-Domain 2 (D2) family, and Lon family. Apart from clade-specific features, these AAA+ families also exhibit distinct complex architectures (Sauer and Baker, 2011) (Figure 7.1). For example, the protease domain and AAA+ module, which consists of a large and small AAA subdomain, are expressed in the same protein subunit for FtsH and Lon, whereas these two kinds of modules reside in distinct subunits in other protease complexes. p97/Cdc48, HslU, and ClpA/B/C even contain two AAA+ modules per subunit. Recent cryogenic electron microscopy (cryo-EM) studies in conjunction with biochemical experiments have elucidated structures and mechanisms of several protease complexes during the process of unfolding or degrading a substrate protein. In this chapter, we will focus on these latest results and describe the structure, function, and working mechanism of AAA-ATPases in the 26S proteasome.

7.2

AAA+ ATPases IN UBIQUITIN-PROTEASOME SYSTEM

Ubiquitin-proteasome system (UPS) and autophagy are two major mechanisms for the degradation of a great majority of the intracellular proteins and unnecessary or dysfunctional components

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FIGURE 7.2 Structure and translocation mechanism of 26S proteasome. (a) Cryo-EM structure of substratebound human proteasome in state EB (EMDB ID: 9218; PDB ID: 6MSE). The RPT1 density is omitted to show the substrate density inside the ATPase ring. (b) A close-up view of the quaternary interface around the isopeptide bond between substrate and ubiquitin. (c) Architecture of pore loop staircase interacting with the substrate. Aromatic residues in pore-1 loops are labelled. (d) Molecular model of RPT5 in state EB, with ATP bound and substrate engaged. (e) Schematic of mechanical substrate translocation of proteasomal ATPases. Synchronization of nucleotide processing in three adjacent ATPases (left) causes differential vertical rigidbody rotations in each substrate-engaged ATPase that cooperatively transfer the substrate (right). (Reproduced from Dong et al., 2019.)

in eukaryotic cells (Ciechanover, 2005; Finley and Prado, 2019; Mayer, 2000). Proteins covalently modified by polyubiquitin chains via a cascade of E1, E2, and E3 enzymes are selectively targeted and destructed by the 26S proteasome, a 2.5-megadalton proteolytic molecular machine equipped with AAA ATPases. As the endpoint of the UPS, the 26S proteasome is the most sophisticated protease complex known, ubiquitously found in all eukaryotes (Darwin, 2009; Muller and Weber-Ban, 2019). There have been some excellent reviews covering proteasome structure and function (Bard et al., 2018; Collins and Goldberg, 2017; Finley et al., 2016; Finley and Prado, 2019; Mao, 2021), assembly (Budenholzer et al., 2017; Rousseau and Bertolotti, 2018), ubiquitin recognition (Kwon and Ciechanover, 2017; Saeki, 2017; Yu and Matouschek, 2017), and proteasomal deubiquitinating enzymes (de Poot et al., 2017). The 26S proteasome holoenzyme assembles from one CP and two RPs capping both sides of the CP cylinder (Figure 7.2a). The CP, also known as 20S proteasome, is composed of distinct α-type and β-type subunits that are stacked into a barrel-like α7β7β7α7 assembly. The RP, also known as 19S or PA700 complex, is the most commonly observed proteasome activator whose assembly and function are dependent on ATP. Other ATP-independent activators, such as 11S (PA28) and Blm10 (PA200), can also be associated with the CP to activate the proteasome holoenzyme (Stadtmueller and Hill, 2011). The RP is structurally composed of a lid and a base subcomplex. The lid subcomplex comprises nine RPN (regulatory particle non-ATPase) subunits (RPN3, RPN5, RPN6, RPN7, RPN8, RPN9, RPN11, RPN12, and Sem1/Dss1). The base subcomplex consists of RPN1, RPN2, RPN13 and six paralogous, distinct RPT (regulatory particle ATPase) subunits (RPT1-RPT6) from the classic AAA family. Another RP subunit RPN10 interacts with both the base and lid and was previously considered to be part of the base. Each RPT subunit consists of an N-terminal helical domain, which dimerizes into a coiled coil (CC) between adjacent subunits (RPT1/RPT2, RPT6/RPT3,

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and RPT4/RPT5), an oligonucleotide- and oligosaccharide-binding (OB) domain, and a C-terminal AAA domain, which comprises a large and small AAA subdomain (Figure 7.2d). Like other AAA+ ATPases, the AAA domain of RPT contains highly conserved motifs, including Walker A, Walker B, sensor 1, arginine finger (R-finger), sensor 2, and pore-1/2 loops (Erzberger and Berger, 2006; Hanson and Whiteheart, 2005; Iyer et al., 2004; Lupas and Martin, 2002; Ogura and Wilkinson, 2001; Sauer and Baker, 2011). The pore loops interact with substrates directly and form a central translocation channel in a right-handed spiral staircase arrangement. The nucleotide-binding pocket of each RPT subunit is surrounded by Walker A, Walker B, sensor 1, and sensor 2 from one “cis” RPT subunit, and two R-fingers from the large AAA subdomain of the clockwise adjacent RPT subunit, functioning in “trans”, allowing for allosteric communication between adjacent subunits (Wendler et al., 2012). With these structural motifs, coordinated ATP binding and hydrolysis around the ATPase ring drive conformational changes of the ATPases, converting the chemical energy of ATP hydrolysis into mechanical work of substrate translocation through the axial channel. Mutagenesis experiments have found that each of the six ATPases exhibited functional asymmetry in substrate degradation although they all share those conserved motifs (Beckwith et al., 2013; Erales et al., 2012; Zhang et al., 2022). Recognition of a ubiquitylated substrate is mediated by the ubiquitin receptors, including RPN1 (Shi et al., 2016), RPN10 (Deveraux et al., 1994; van Nocker et al., 1996), and RPN13 (Husnjak et al., 2008; Schreiner et al., 2008). After substrate recruitment, a flexible initiation region of the substrate is then captured by the pore loops of the RPT subunits (Bard et al., 2019; Dong et al., 2019; Prakash et al., 2004; Yu and Matouschek, 2017). To allow subsequent degradation, conjugated ubiquitin chains are removed by either the stoichiometric deubiquitylating enzyme (DUB) subunit RPN11 (Dong et al., 2019; Verma et al., 2002; Worden et al., 2017; Yao and Cohen, 2002) or the reversibly associated DUBs like Ubp6/USP14 (Lee et al., 2016) and Uch37/UCHL5 (Vander Linden et al., 2015). The globular domains of an engaged substrate are then mechanically unfolded and translocated through the narrow axial channel of the heterohexameric ATPase ring made of six distinct RPT subunits. Meanwhile, the central entry port of the CP proteolytic chamber (a.k.a. CP gate) remains closed in the resting state (Chen et al., 2016; Huang et al., 2016; Schweitzer et al., 2016). Opening of the CP gate is triggered by docking of five C-terminal tails of all RPT subunits except RPT4 into the inter-subunit surface pockets of the α-ring (a.k.a. α-pockets), which are formed between adjacent α-subunits (Chen et al., 2016; Dong et al., 2019; Eisele et al., 2018; Wehmer et al., 2017; Zhu et al., 2018; Rabl et al., 2008; Smith et al., 2007). Structural studies on the human 26S proteasome have established that the insertion of the C-termini of RPT3 and RPT5 into α-pockets, which contain conserved hydrophobic-Tyr-X (HbYX) motifs, ensures RP’s association with CP but is not sufficient for CP gate opening (Chen et al., 2016; Huang et al., 2016; Schweitzer et al., 2016). By contrast, the insertion of Rpt2 C-terminus also contributes to the yeast 26S proteasome assembly in addition to those of Rpt3 and Rpt5 (Ding et al., 2017; Eisele et al., 2018; Wehmer et al., 2017). In both human and yeast proteasomes, CP gate opening was observed when the C-termini of all RPT subunits except RPT4 were engaged with the α-pockets (Chen et al., 2016; Zhu et al., 2018; Eisele et al., 2018). Human p97 or valosin-containing protein (VCP) in higher eukaryotes, and its yeast ortholog Cdc48, remodel or reprocess ubiquitinated substrates for ubiquitin-dependent degradation (Meyer et al., 2012; Stolz et al., 2011; van den Boom and Meyer, 2018; Xia et al., 2016), and play an important role in the UPS pathway (Godderz et al., 2015), especially in endoplasmic reticulum (ER)-associated protein degradation (ERAD) (Christianson and Ye, 2014; Stein et al., 2014; Wolf and Stolz, 2012; Wu and Rapoport, 2018) and outer mitochondrial membrane-associated degradation (OMMAD) (Heo et al., 2010; Taylor and Rutter, 2011; Xu et al., 2011). It has been suggested that archaeal Cdc48 can artificially assemble with 20S proteasome in vitro through crosslinking (Barthelme et al., 2014). Since the 26S proteasome requires an unstructured polypeptide segment in its substrate to initiate processing (Fishbain et al., 2011; Inobe et al., 2011; Prakash et al., 2004), p97/Cdc48 can act upstream of the proteasome when the substrate is located in membranes or well folded without a flexible initiation region (Beskow et al., 2009; Olszewski et al., 2019). p97/Cdc48 can partially

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or completely unfold the substrate and transfer it to the 26S proteasome for degradation with the assistance of the shuttling factors Rad23 or Dsk2 (Baek et al., 2011; Itakura et al., 2016; Richly et al., 2005), or directly into the 20S complex (Barthelme and Sauer, 2012, 2013). Furthermore, p97/Cdc48 is also involved in other cellular processes, including autophagy (Ju et  al., 2009; Papadopoulos et al., 2017), ribosomal quality control (Brandman et al., 2012; Verma et al., 2013), extraction of chromatin-bound proteins (Dantuma and Hoppe, 2012; Franz et al., 2016; Ramadan et al., 2007), membrane fusion, and vesicular trafficking (Bug and Meyer, 2012; Patel et al., 1998). It is associated with several diseases (Chapman et al., 2011; Kimonis et al., 2008; Tang and Xia, 2016) and has also been identified as a promising target for anticancer drugs due to its crucial roles in protein quality control, homeostasis, and cell viability (Anderson et al., 2015; Magnaghi et al., 2013; Skrott et al., 2017; Vekaria et al., 2016). Similar to the proteasomal RP, AAA+ ATPases in the p97/Cdc48 complex can unfold and translocate a substrate through its central pore. In addition to an N-terminal (N) domain and a flexible C-terminal tail, a p97/Cdc48 monomer encompasses two tandem ATPase domains (D1 and D2), each forming a ring-like homohexamer (Figure 7.1). Both D1 and D2 are homologous to single AAA domain of PAN and proteasomal RPT subunits, hosting a nucleotide-binding pocket and pore-1/2 loops that can interact with substrates (Erzberger and Berger, 2006; Hanson and Whiteheart, 2005; Iyer et al., 2004; Lupas and Martin, 2002; Ogura and Wilkinson, 2001; Pamnani et al., 1997; Sauer and Baker, 2011). Thus, one p97/Cdc48 complex possesses twelve ATP-binding sites in total. The N domains are not fixed with respect to the double ring (Davies et al., 2008; Zhang et al., 2000). Upon ATP binding to the D1 ATPases, N domains are displaced from a “down-conformation” coplanar with the D1 ring to an “up-conformation” above the D1 plane (Banerjee et al., 2016; Tang et al., 2010). To engage a substrate, p97/Cdc48 often needs the assistance of various cofactors such as Ufd1/Npl4 heterodimer, which usually bind to the N domains or the C-termini for regulating substrate recognition (Buchberger et al., 2015; Hanzelmann and Schindelin, 2017). These cofactors endow p97/Cdc48 with the substrate specificity and pathway selectivity.

7.3

CONFORMATIONAL CHANGES OF AAA ATPases IN THE 26S PROTEASOME

In contrast to high stability of the CP (Groll et al., 1997; Lowe et al., 1995), the RP and particularly the AAA-ATPases module exist as a highly dynamic component, sampling an expanded conformational landscape (Chen et al., 2016; de la Pena et al., 2018; Dong et al., 2019; Lu et al., 2017; Unverdorben et al., 2014; Zhu et al., 2018). Recent studies of single-particle cryo-EM and cryoelectron tomography (cryo-ET) suggested that the proteasome holoenzymes in cells mostly stay in a basal resting state in the presence of ATP and in the absence of substrate, and spontaneously sample several alternative conformations (Asano et al., 2015; Chen et al., 2016; Haselbach et al., 2017; Unverdorben et al., 2014). It has been found that the conformational distributions of the 26S proteasome can be modified by using hydrolysis-inactivated Walker-B mutations (Eisele et al., 2018), deactivating certain subunit with mutations or inhibitors (Haselbach et al., 2017; Matyskiela et al., 2013), and replacing ATP with slowly hydrolyzed ATPγS or nonhydrolyzable ATP analogs (Ding et al., 2017; Sledz et al., 2013; Unverdorben et al., 2014; Wehmer et al., 2017; Zhu et al., 2018). At least six distinct conformations of both human and yeast proteasomes without any substrate bound have been observed (Zhu et al., 2018; Bard et al., 2018; Mao, 2021). Since the proteolysis process was absent, the observed conformational changes mainly reflect an idle ATPase motor with no external mechanical work output. Recent cryo-EM studies of substrate-engaged 26S proteasomes offered the first high-resolution views of dynamic substrate-proteasome interactions and insights into the inner workings of this macromolecular machine (de la Pena et al., 2018; Dong et al., 2019; Wu et al., 2021; Zhang et al., 2022). Unlike several studies that completely replace ATP with ATPγS or nucleotide analogs

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(Eisele et al., 2018; Sledz et al., 2013; Wehmer et al., 2017; Yu et al., 2018; Zhu et al., 2018), Dong et al. devised a novel time-dependent “nucleotide-substitution” strategy, which first primes the substrateengaged proteasome with ATP, then dilutes ATP with ATPγS within 30 seconds after the initial phase of substrate engagement with the human 26S proteasome to quench the hydrolytic activity of AAA+ ATPases (Dong et al., 2019). This approach maximizes the conformational diversity and heterogeneity of 26S proteasome, allowing critical functional intermediates to be captured before the completion of degradation reactions. To compensate the complexity of cryo-EM analysis conferred by the extreme conformational heterogeneity in this case, the researchers collected a considerably large cryo-EM dataset, extensively used the latest-developed machine-learning tools in data clustering (Wang et al., 2019; Wu et al., 2017; Xu et al., 2016; Zhu et al., 2017) and eventually sorted out seven conformations of the substrate-bound human 26S proteasome at 2.8-3.6 Å resolution (Dong et al., 2019). Application of deep-learning-enhanced 3D classification further expanded the differentiated states into 64 conformers, with majority reaching near-atomic resolution (Wu et al., 2021; Wu et al., 2022). Together, these structural snapshots depict a spatiotemporal continuum of functional proteasome conformations during polyubiquitylated-substrate degradation, shedding light on the complete cycle of substrate processing by the human proteasome, from initial ubiquitin and UBL recognition (Dong et al., 2019; ;Wu et al., 2021; Zhang et al., 2022), RPN11 and USP14-catalyzed deubiquitylation (Dong et al., 2019; Zhang et al., 2022), and translocation initiation (Dong et al., 2019; Wu et al., 2021), to processive substrate degradation (Dong et al., 2019; Zhang et al., 2022). The conformational changes of RPT subunits in the proteasome appear to be strongly coupled to all major steps of substrate processing and play an important role in dynamic regulation of the proteasome function (Dong et al., 2019; Zhang et al., 2022). Although the overall structural relationships between the RP and CP and between the lid and base seem to be highly consistent among the 26S proteasome conformations with or without substrates, one of the key conformations termed state EB, representing the human proteasome under RPN11-catalyzed deubiquitylation, appears to be missing in all previous studies except for one (Dong et al., 2019). The proteasome structure of state EB reveals an unexpected quaternary subcomplex involving RPN11, RPN8, and RPT5 (Figure 7.2b) (Dong et al., 2019), which was also observed in states EA2.1UBL and EA2.2UBL in the presence of USP14 (Zhang et al., 2022). Around the scissile isopeptide bond between the RPN11-bound ubiquitin and the substrate lysine, a ternary interface is formed between RPN11, RPN8, and the N-loop of RPT5, which emanates from the top of its OB domain to efficiently carry out the deubiquitylation mission (Figure 7.2b) (Dong et al., 2019). The N-terminal CC domains of RPT subunits, contacting with the lid subcomplex, allosterically regulate ATPase activity in a long-range fashion and contribute to conformational switch of the holoenzyme (Dong et al., 2019; Snoberger et al., 2018). The RP undergoes dramatic rotation (30°–40°) and translation above the CP during the transition of CP gate from the closed to open state. The revelation of state EB fills up a fundamentally critical intermediate missed in all early studies (Chen et al., 2016; Huang et al., 2016; Schweitzer et al., 2016; Zhu et al., 2018), allowing the observation of stepwise activation of the CP by the insertion of RPT C-tails into the α-pockets one at a time (Dong et al., 2019). During the process of CP gate opening, the relative position of the catalytic site of DUB RPN11, the translocation channel of ATPase ring, and the CP gate are aligned coaxially step-by-step (de la Pena et al., 2018; Dong et al., 2019; Matyskiela et al., 2013; Sledz et al., 2013), so that the substrate polypeptide can be threaded processively into the proteolytic CP chamber.

7.4 SUBSTRATE INTERACTIONS COUPLED WITH ATP HYDROLYSIS In each ATPase subunit, the ATP hydrolytic cycle converts chemical energy into mechanical work on the substrate bound to a staircase of pore loops (Erzberger and Berger, 2006; Hanson and Whiteheart, 2005; Iyer et al., 2004; Lupas and Martin, 2002; Ogura and Wilkinson, 2001; Sauer and Baker, 2011). The quaternary architecture of the pore loops narrowing the inner channel appears to be conserved among different translocase systems (Beckwith et al., 2013; Erales et al., 2012; Hinnerwisch et al., 2005; Martin et al., 2008) (Figure 7.2c). Substrate-contacting pore-1 loops

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of four or five ATPase subunits are approximately evenly distributed along the unfolded substrate polypeptide and form an architecture of spiral staircase, with two adjacent pore-1 loops spanning two amino acid residues along the substrate, presumably corresponding to one step in translocation driven by the hydrolysis of single ATP molecule (Dong et al., 2019). This “two-residue spacing” of substrate-contacting pore-1 loops appears to be a key structural “rule” that is highly conserved among many AAA+ ATPases, including 26S proteasome (de la Pena et al., 2018; Dong et al., 2019; Wu et al., 2021; Zhang et al., 2022), p97/Cdc48 (Olszewski et al., 2019; Cooney et al., 2019; Twomey et al., 2019; Ripstein et al., 2017; Pan et al., 2021; Xu et al., 2022), FtsH-like AAA proteases (Puchades et al., 2019; Puchades et al., 2017), and Hsp104 disaggregase (Gates et al., 2017). It is expected that a conserved mechanism may underlie the force generation by intercalated stacking interactions between the pore-1 loop’s aromatic residues and the substrate sidechains. The pore-2 loops form a similar but shorter staircase underneath the pore-1 loops, supporting the opposite side of the substrate through charged acidic residues (Chen et al., 2016; de la Pena et al., 2018; Dong et al., 2019; Zhu et al., 2018; Zhang et al., 2022). The nucleotide-binding pocket at the Walker B motif is located near a short-loop hinge connecting the large and small AAA subdomains (Figure 7.2d). ATP-binding pocket is embraced by both AAA subdomains. Thus, the occupancy and type of nucleotide at this pocket determines the hinge configuration and ATP binding locks the AAA domain into a single rigid body. When ATP is hydrolyzed into ADP and γ-phosphate is subsequently released, the corresponding ATPase subunit undergoes an outward flipping of 30°–40°, resulting in disengagement of the ATPase from the substrate (Figure 7.2e). Three states of nucleotide-binding pocket were observed: ATP-bound, ADP-bound, and apo-like state in which only a very weak or partial density is exhibited inside its nucleotidebinding pocket. Systematic structural alignment of ATPases among different states showed that ATP binding or ADP release leads to a hinge-like rotation of 15°–25° between its small and large AAA subdomains (Dong et al., 2019). By contrast, release of γ-phosphate after ATP hydrolysis appears to be insufficient to immediately trigger inter-subdomain motion within the AAA domain. Under this circumstance, the whole AAA domain could still rotate as a rigid body upon inter-subunit interactions. The subsequent release of ADP liberates the free energy and triggers a hinge-like rotation between the large and small AAA subdomains that disengages the corresponding ATPase subunit from the substrate, and spreads the kinetic energy out to drive rigid-body rotation of four or five substrate-bound ATPase subunits of the holoenzyme, propelling the substrate forward (de la Pena et al., 2018; Dong et al., 2019). These structural findings are most compatible with a sequential ATP hydrolysis model than with a random one in the ATPase hexameric ring (Figures 7.2e and 7.3). One of the notable features in this model is that not all ATPase subunits contact with the substrate simultaneously; instead, at least one subunit is disengaged from the substrate upon ADP release. This appears to be a common feature observed in most substrate-bound ATPase hexamer structures (Alfieri et al., 2018; Deville et al., 2017; Gates et al., 2017; Han et al., 2017; Monroe et al., 2017; Puchades et al., 2019; Puchades et al., 2017; Ripstein et al., 2017; Thomsen and Berger, 2009).

7.5

THREE MODES OF COORDINATE ATP HYDROLYSIS REGULATE INTERMEDIATE FUNCTIONAL STEPS

The ATPase ring in the 26S proteasome bears a greater degree of inter-subcomplex interactions than most of other AAA+ ATPase complexes. On one side, the ATPase ring forms extensive intersubunit interfaces with the lid subcomplex and the RPN subunits in the base. On the other side, the ATPase ring forms a multivalent, highly dynamic interface with the α-ring in the CP, mostly via the C-terminal tails of five RPT subunits. The structural complexity might well have been evolved to accommodate the functional complexity of the proteasome in ubiquitin recognition, selective substrate engagement and timed deubiquitylation, substrate unfolding and processive translocation. Unexpectedly, three distinct modes of coordinated ATP hydrolysis in the proteasomal ATPase ring

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FIGURE 7.3 Schematic of coordinated ATP hydrolysis and nucleotide exchange observed in seven states in the substrate-bound human 26S proteasome (Dong et al., 2019). Three principal modes are depicted here, respectively, featuring hydrolytic events in two oppositely positioned ATPases, in two adjacent ATPases and in one ATPase at a time. (Reproduced from ref. Dong et al., 2019.)

have been discovered to regulate these key functional steps of proteasome (Figure 7.3) (Dong et al., 2019). The ability of functioning in multiple modes by the AAA-ATPase motor suggests the existence of multiple pathways of conformational transitions induced by coordinated ATP hydrolysis and inter-subcomplex interactions (Mao, 2021; Zhang et al., 2022).

7.5.1

Mode 1 Regulates ubiquitin ReCognition, initial substRate engageMent, and deubiquitylation

Mode 1 features coordinated ATP hydrolysis in a pair of oppositely positioned ATPases and was observed in states EA1, EA2, and EB of the human 26S proteasome corresponding to the intermediate steps of initial ubiquitin recognition and deubiquitylation (Peth et al., 2010; Worden et al., 2017). This mode has been observed in both the presence and absence of USP14 (Dong et al., 2019; Zhang et al., 2022). Before the proteasome in state EB gets ready to remove the ubiquitin chain from substrate with the DUB RPN11, the ADP bound to RPT6 in state EA is released. Meanwhile, the ATP molecules in both RPT2 and its opposite subunit RPT4 get hydrolyzed. These events drive an outward rotation and partial refolding of RPT6 and an iris-like movement in the AAA ring that opens its axial channel for initial substrate insertion into the AAA ring. Similar mode of coordinated ATP hydrolysis was also observed in the crystal structure of substrate-free hexameric ClpX protease (Glynn et al., 2009), which drives rather different conformational changes in the ATPase ring, compared to those in the 26S proteasome.

7.5.2

Mode 2 Regulates Cp gating, ubiquitin Release, and initiation oF substRate tRansloCation

Mode 2 features coordinated ATP hydrolysis in at least two adjacent ATPases and was observed in states EC1 and EC2 of the human 26S proteasome corresponding to the intermediate steps of initiation

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of substrate translocation while preparing for CP gate opening and activation in the absence of USP14. A key structural feature of this mode is the simultaneous disengagement of two adjacent ATPases from the substrate. After RPN11-catalyzed deubiquitylation, the proteasome is about to initiate substrate translocation and prepare for allosteric regulation of the CP gate opening. To this end, ATP molecules in two adjacent subunits, RPT1 and RPT5, are hydrolyzed, with RPT1 and its clockwise neighbor, RPT2, disengaged from the substrate. As RPT6 rebinds ATP and returns to the top of the substrate-bound poor-loop staircase, the conformational changes from state EB to EC result in a one-step forward translocation of the substrate by two residues. However, during the following EC2-to-ED1 transition, both the substrate-disengaged RPT1 and RPT2 need to bind ATP and return to the top of the substrate-bound pore loop staircase, and the RPT5 is about to release its ADP, which together drive a two-step forward translocation of the substrate. The second half of this transition has been resolved in six high-resolution conformations that allowed visualization of single-nucleotide exchange dynamics in RPT1 (Wu et al., 2021). Recent cryo-EM structures of the D2 ATPase ring of yeast Cdc48 and of FtsH-like mitochondrial protease AFG3L2 exhibited a substrate-bound ATPase architecture similar to Mode 2 of the 26S proteasome (Puchades et al., 2019; Twomey et al., 2019). Surprisingly, the USP14-regulated human proteasome appears to either considerably attenuate the probability of Mode 2 or forgo this conformational transition pathway (Zhang et al., 2022).

7.5.3

Mode 3 Regulates pRoCessive substRate unFolding, tRansloCation, and degRadation

Mode 3 features coordinated ATP hydrolysis in only one ATPase at a time and was observed in states ED1 and ED2 of the human 26S proteasome corresponding to the intermediate steps of processive substrate unfolding and translocation. When the pore-1 loop of an RPT subunit reaches the CP-proximal position at the bottom of the substrate-bound pore-loop staircase, this RPT subunit is always ADP-bound. Then, the ADP molecule is released from the binding pocket, and the subsequent hinge-like rotation between the small and large AAA subdomains disengages the pore loop from the substrate and flips this RPT subunit outwards away from the ATPase ring. At the same time, its counterclockwise adjacent ADP-bound RPT subunit is pushed to the bottom of the substrate-bound pore-loop staircase; its clockwise adjacent RPT subunit, which was an apo-like detached seam earlier, now acquires a new ATP and reengages with the substrate at the top of the staircase via a hinge-like rotation. Concomitantly with these concerted motions, the other three substrate-engaged subunits are mostly ATP-bound and rotate downwards approximately as a rigid body in concert driven by the conformational changes in the RPT subunits undergoing nucleotide exchange. In the study on the PAN (proteasome-activating nucleotidase)-activated proteasome, an archaea homolog of the 26S proteasome, five distinct conformations in the PAN ATPase ring observed in the absence of substrate all have only one ATPase disengaged from the rest of the ATPase ring, a key feature consistent with Mode 3 (Majumder et al., 2019). Similar substrate-bound AAA+ ATPase ring conformations were also observed in p97/Cdc48 (Cooney et al., 2019; Twomey et al., 2019; Pan et al., 2021; Xu et al., 2022). Although only one step of substrate translocation has been observed in Mode 3 in the absence of USP14 (de la Pena et al., 2018; Dong et al., 2019), a near-complete cycle of coordinated ATP hydrolysis around the ATPase ring with an open CP gate during processive translocation was observed in six distinct ATPase ring conformations in the presence of USP14 (Zhang et al., 2022). These structural observations instead suggest that asymmetric ATP hydrolysis around the ATPase ring is coupled to metastability of intermediate conformations with two ATPase subunits disengaged from substrate during processive substrate translocation (Zhang et al., 2022; Fang et al., 2022), which is deviated from initial hypothesis of a rigorous Mode-3 action (Dong et al., 2019).

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Biomotors and their Nanobiotechnology Applications

EVIDENCE FOR A SEQUENTIAL HAND-OVER-HAND MODEL

While most other substrate-bound AAA+ ATPase structures were solved in only one or few conformations at high resolution per biochemical condition (Cooney et al., 2019; Monroe et al., 2017; Puchades et al., 2019; Puchades et al., 2017; Ripstein et al., 2017; de la Pena et al., 2018; Majumder et al., 2019; Pan et al., 2021; Xu et al., 2022), tens of coexisting conformations of the proteasome at atomic details have provided unprecedented insights into both single-nucleotide-exchange dynamics of the ATPase (Wu et al., 2021) and the complete around-ring cycle of ATP hydrolysis (Dong et  al., 2019; Zhang et al., 2022). Notably, the revelation of sequential hand-over-hand action of asymmetric, coordinated ATP hydrolysis around the complete ring of the ATPase motor in the presence of an open CP gate was recapitulated in the computational simulations of substrate translocation in the human proteasomal AAA-ATPase motor (Saha and Warshel, 2021; Fang et al., 2022) and further substantiated in the experimental studies of USP14-regulated human proteasome in the act of substrate processing by using time-resolved cryo-EM (Zhang et al., 2022). These investigations together further verified the time sequence of the corresponding states along the paths of chemical reactions. The design of AAA+ ATPase systems should be versatile enough to allow for the coexistence of multiple pathways of coordinated ATP hydrolysis. The existence of a rigorously sequential Mode-3 hydrolysis does not necessarily exclude the possibility of less sequential, more stochastic ATP hydrolysis or with mixed modes that are compatible with the hand-over-hand mechanism in general. Indeed, USP14 activation by the proteasome reprograms the ATPase motor and appears to stabilize substrate-bound conformations with two ATPase subunits disengaged from the substrate during processive substrate translocation (Zhang et al., 2022). In addition, recent cryo-EM structure of the yeast Cdc48 in complex with its heterodimeric cofactor Ufd1-Npl4 and a polyubiquitinated substrate reveal a nearly planar D1 ATPase ring and a spiral-shaped D2 ATPase ring in Mode-2like conformation (Twomey et al., 2019). However, the use of ADP/BeFx resulted in a spiral D1 ring with the D2 ring turning into Mode-3-like conformation. Similarly, cryo-EM structures of two FtsH-like mitochondrial proteases, the engineered soluble Yme1 and AFG3L2 complexes, revealed conformations of substrate-bound ATPase ring highly resembling Modes 3 and 2 conformations of the 26S proteasome, respectively (Puchades et al., 2019; Puchades et al., 2017). Thus, the fate of the ATP hydrolysis pathway in the hexameric ATPase ring is likely energetically dependent on their interactions with specific substrates, regulatory subunits, chaperones, or cofactors, which deserves further in-depth investigation.

7.7

CONCLUDING REMARKS

In this chapter, we summarize the recent significant discoveries of molecular mechanism of proteasomal AAA-ATPase motor in the 26S proteasome that is representative of the classic clade of AAA+ ATPase superfamily and has far-reaching implications in health and disease. A sequential hand-over-hand mechanism of substrate translocation driven by coordinated ATP hydrolysis unidirectionally around the ATPase ring appears to be the favored model for substrate translocation through many ring-like ATPase hexamers (de la Pena et al., 2018; Dong et al., 2019; Majumder et al., 2019; Puchades et al., 2017; Pan et al., 2021; Xu et al., 2022; Saha and Warshel, 2021; Fang et al., 2022). To date, the greatest number of distinct conformations of the same ATPase motor analyzed (more than 30) in the context of human proteasome holoenzyme complex have provided the most extensive picture of functional AAA+ ATPase dynamics at the atomic level (Chen et al., 2016; Dong et al., 2019; Zhu et al., 2018; Wu et al., 2021; Zhang et al., 2022). Multiple pathways of coordinated ATP hydrolysis around the ATPase ring were found to be associated with functional intermediate steps of substrate processing in the human proteasome (Dong et al., 2019; Zhang et al., 2022). Together, these studies not only revealed highly conserved aspects of substrate translocation mechanisms in various AAA+ ATPase hexamers (Glynn et al., 2009; Puchades et al., 2019;

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Cooney et al., 2019; Twomey et al., 2019; de la Pena et al., 2018; Deville et al., 2019; Gao et al., 2019; Puchades et al., 2017; Ripstein et al., 2020; Yu et al., 2018; Pan et al., 2021; Xu et al., 2022; Wald et al., 2022) but also characterized the parallel pathways of conformational dynamics that are uniquely used by the 26S proteasome (Dong et al., 2019; Wu et al., 2021; Saha and Warshel, 2021; Zhang et al., 2022; Fang et al., 2022). Future investigations into the detailed mechanisms of proteasome regulation by other important factors as well as an integrative, quantitative, self-consistent model of functional complex dynamics at both molecular and systems levels will hopefully bring us to another whole new level of understanding of the ATPase motor complexity.

FUNDING This work was partly funded by Natural Science Foundation of Beijing Municipality, Grant Number Z180016, and National Natural Science Foundation of China, Grant Number 12125401 and 1177402.

ACKNOWLEDGMENTS The authors thank all members of the Mao Laboratory for helpful discussions and particularly Shitao Zou for a proofreading of the manuscript; and Drs. Marc Kirschner, Ying Lu, Daniel Finley, Peixuan Guo, and Alfred Goldberg for constructive, insightful discussions relevant to this work. We apologize for not including many relevant excellent literatures due to our current focus and scope of this review and our limitation in referencing efforts.

CONFLICTS OF INTEREST The authors declare no conflict of interest.

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Unverdorben, P., Beck, F., Sledz, P., Schweitzer, A., Pfeifer, G., Plitzko, J.M., Baumeister, W., and Forster, F. (2014). Deep classification of a large cryo-EM dataset defines the conformational landscape of the 26S proteasome. Proc Natl Acad Sci U S A 111, 5544–5549. Vale, R.D. (2000). AAA proteins. Lords of the ring. J Cell Biol 150, F13–F19. van den Boom, J., and Meyer, H. (2018). VCP/p97-mediated unfolding as a principle in protein homeostasis and signaling. Mol Cell 69, 182–194. van Nocker, S., Sadis, S., Rubin, D.M., Glickman, M., Fu, H., Coux, O., Wefes, I., Finley, D., and Vierstra, R.D. (1996). The multiubiquitin-chain-binding protein Mcb1 is a component of the 26S proteasome in Saccharomyces cerevisiae and plays a nonessential, substrate-specific role in protein turnover. Mol Cell Biol 16, 6020–6028. Vander Linden, R.T., Hemmis, C.W., Schmitt, B., Ndoja, A., Whitby, F.G., Robinson, H., Cohen, R.E., Yao, T., and Hill, C.P. (2015). Structural basis for the activation and inhibition of the UCH37 deubiquitylase. Mol Cell 57, 901–911. Vekaria, P.H., Home, T., Weir, S., Schoenen, F.J., and Rao, R. (2016). Targeting p97 to disrupt protein homeostasis in cancer. Front Oncol 6, 181. Verma, R., Aravind, L., Oania, R., McDonald, W.H., Yates, J.R., 3rd, Koonin, E.V., and Deshaies, R.J. (2002). Role of Rpn11 metalloprotease in deubiquitination and degradation by the 26S proteasome. Science 298, 611–615. Verma, R., Oania, R.S., Kolawa, N.J., and Deshaies, R.J. (2013). Cdc48/p97 promotes degradation of aberrant nascent polypeptides bound to the ribosome. Elife 2, e00308. Wald, J., Fahrenkamp, D., Goessweiner-Mohr, N., Lugmayr, W., Ciccarelli, L., Vesper, O., and Marlovits, T.C. (2022). The Mechanism of AAA+ ATPase-mediated RuvAB-Holliday junction branch migration. Nature 609, 630–639. Wang, J., Hartling, J.A., and Flanagan, J.M. (1997). The structure of ClpP at 2.3 Å resolution suggests a model for ATP-dependent proteolysis. Cell 91, 447–456. Wang, W.L., Yu, Z., Castillo-Menendez, L.R., Sodroski, J., and Mao, Y. (2019). Robustness of signal detection in cryo-electron microscopy via a bi-objective-function approach. BMC Bioinform 20, 169. Wehmer, M., Rudack, T., Beck, F., Aufderheide, A., Pfeifer, G., Plitzko, J.M., Forster, F., Schulten, K., Baumeister, W., and Sakata, E. (2017). Structural insights into the functional cycle of the ATPase module of the 26S proteasome. Proc Natl Acad Sci U S A 114, 1305–1310. Wendler, P., Ciniawsky, S., Kock, M., and Kube, S. (2012). Structure and function of the AAA+ nucleotide binding pocket. Biochim Biophys Acta 1823, 2–14. Wolf, D.H., and Stolz, A. (2012). The Cdc48 machine in endoplasmic reticulum associated protein degradation. Biochim Biophys Acta 1823, 117–124. Worden, E.J., Dong, K.C., and Martin, A. (2017). An AAA motor-driven mechanical switch in Rpn11 controls deubiquitination at the 26S proteasome. Mol Cell 67, 799–811. Wu, J., Ma, Y.B., Congdon, C., Brett, B., Chen, S., Xu, Y., Ouyang, Q., and Mao, Y. (2017). Massively parallel unsupervised single-particle cryo-EM data clustering via statistical manifold learning. PLoS One 12, e0182130. Wu, X., and Rapoport, T.A. (2018). Mechanistic insights into ER-associated protein degradation. Curr Opin Cell Biol 53, 22–28. Wu, Z., ., Chen, E., Zhang, S., Wang, W.L., Dong, Y., and Mao, Y. (2021). Hidden dynamics of human proteasome autoregulation during protein degradation. BioRxiv https://doi.org/10.1101/2020.12.22.423932. Wu, Z., ., Chen, E., Zhang, S., Ma, Y., and Mao, Y. (2022). Visualizing conformational space of functional biomolecular complexes by deep manifold learning. Int J Mol Sci 23, 8872. Xia, D., Tang, W.K., and Ye, Y. (2016). Structure and function of the AAA+ ATPase p97/Cdc48p. Gene 583, 64–77. Xu, S., Peng, G., Wang, Y., Fang, S., and Karbowski, M. (2011). The AAA-ATPase p97 is essential for outer mitochondrial membrane protein turnover. Mol Biol Cell 22, 291–300. Xu, Y., Han, H., Cooney, I., Guo, Y., Moran, N.G., Zuniga, N.R., Price, J.C., Hill, C.P., and Shen, P.S. (2022). Active conformation of the p97-p47 unfoldase complex. Nat Commun 13, 2640. Xu, Y., Wu, J., Yin, C.C., and Mao, Y. (2016). Unsupervised Cryo-EM data clustering through adaptively constrained K-means algorithm. PLoS One 11, e0167765. Yao, T., and Cohen, R.E. (2002). A cryptic protease couples deubiquitination and degradation by the proteasome. Nature 419, 403–407. Yu, H., Lupoli, T.J., Kovach, A., Meng, X., Zhao, G., Nathan, C.F., and Li, H. (2018). ATP hydrolysis-coupled peptide translocation mechanism of Mycobacterium tuberculosis ClpB. Proc Natl Acad Sci U S A 115, E9560–E9569.

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Yu, H., and Matouschek, A. (2017). Recognition of client proteins by the proteasome. Annu Rev Biophys 46, 149–173. Zhang, S., Zou, S., Yin, D., Zhao, L., Finley, D., Wu, Z., and Mao, Y. (2022). Structure USP14-regulated allostery of the human proteasome by time-resolved cryo-EM. Nature 605, 567–574. Zhang, X., Shaw, A., Bates, P.A., Newman, R.H., Gowen, B., Orlova, E., Gorman, M.A., Kondo, H., Dokurno, P., Lally, J., et al. (2000). Structure of the AAA ATPase p97. Mol Cell 6, 1473–1484. Zhu, Y., Ouyang, Q., and Mao, Y. (2017). A deep convolutional neural network approach to single-particle recognition in cryo-electron microscopy. BMC Bioinform 18, 348. Zhu, Y., Wang, W.L., Yu, D., Ouyang, Q., Lu, Y., and Mao, Y. (2018). Structural mechanism for nucleotidedriven remodeling of the AAA-ATPase unfoldase in the activated human 26S proteasome. Nat Commun 9, 1360.

8

General Mechanism of Biomotors Peixuan Guo and Zhengyi Zhao University of Kentucky

CONTENTS 8.1 Force Generation and Energy Conversion ..............................................................................99 8.2 Motor Subunit Communication ..............................................................................................99 References ...................................................................................................................................... 101

8.1 FORCE GENERATION AND ENERGY CONVERSION In all biomotors, the nucleotide-binding and hydrolysis cycles are coupled to conformational entropy rearrangements of substrate-binding subunits. To date, there are three primary chemomechanical coupling models for biomotors: sequential (individual ATP binding/hydrolysis events proceed sequentially), concerted (all active sites hydrolyze ATP simultaneously), and stochastic (any ATPase site can hydrolyze nucleotide randomly) models. In general, binding of ATP to the disordered ATPase subunit stimulates a conformational change with entropy alteration (De-Donatis et al., 2014) of the ATPase, thus fastening the ATPase at a less random configuration. This new conformation entropy enables the ATPase subunit to bind dsDNA and prime ATP hydrolysis. ATP hydrolysis triggers the second entropic and conformational change, which renders the ATPase into a low affinity for dsDNA and thus pushes the DNA to the next subunit that has already bound to ATP with a high affinity for dsDNA. These continuous actions will promote the movement of the dsDNA around the internal ring.

8.2

MOTOR SUBUNIT COMMUNICATION

The signal transfer modes among motor subunits in different ring translocases are not identical. Sequential action of the phi29 dsDNA packaging motor was originally reported by Chen and Guo (1997) and subsequently confirmed by Bustamante and coworkers (Moffitt et al., 2009). Hill constant determination and binomial distribution of inhibition assay have led to the conclusion that ATPase subunits work sequentially and cooperatively (Schwartz et al., 2013; De-Donatis et al., 2014). This action enables the motor to work continuously without interruption, despite some observable pauses. Extensive studies towards the study of motor subunit communications draw researchers’ attention to the arginine finger motif in the biological motors. Functions of arginine finger motifs slightly vary among different ATPases. The overall structural features of the AAA+ core domains are conserved in all ATPases of the superfamily with a highly conserved arginine residue close to the sensor 2. The helicase superfamily III proteins possess an aberrantly formed α-helical domain; thus, they lack the sensor 2 arginine. The polar residues in sensor 1 were reported to mediate conformational changes, though not all AAA+ proteins possess a polar residue in the sensor 1 position. In the case of p97 D2, the movement of the sensor 1 residue upon nucleotide engagement in the binding pocket will induce displacements at the distal end of the ASCE domain, where the arginine fingers are located. The conserved arginine in the α-helical subdomain was termed sensor 2 residue, which contacts the phosphate groups of ATP and mediates DOI: 10.1201/9780429203367-9

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a conformational change that sequesters the catalytic site from water. Mutations of the sensor 2 residues lead to a loss or decrease of ATP binding and/or ATP hydrolysis (Hanson & Whiteheart, 2005; Chen et al., 2010; Elles & Uhlenbeck, 2008). They also seem to be important for the stability of hexameric complex, since arginine finger mutations in phi29 gp16 ATPase, HslU, p97 VCP, ClpB D1, ClpC D1, and Hsp104 D1 impair oligomer formation even in the presence of ATP (Hanson & Whiteheart, 2005). Hence, the residues not only participate in hydrolysis, but also account for subunit interface contacts and/or the structural integrity of the binding pocket (Hanson & Whiteheart, 2005; Chen et al., 2010; Elles & Uhlenbeck, 2008); in other words, motor subunit communication. Taking the most recent report about phi29 gp16 ATPase as an example, the arginine finger in the ATPase was aligned and mutated (Zhao et al., 2016; Zhao et al., 2017). The arginine finger-free ATPase has shown abolished activities in ATP binding/hydrolysis, nucleotide binding, oligomerization, and in vitro virion assembly. Ultracentrifugation assays and electrophoresis mobility shift assays (EMSA) have demonstrated that arginine finger in gp16 ATPase extends from one subunit to the adjacent one and bridges dimer formation (Zhao et al., 2016; Zhao et al., 2017). Arginine mutants alone could not form dimers, while interactions were observed when they were mixed with either the wild type or other mutants that contained an intact arginine finger, which can provide an arginine residue for dimer formation. Interestingly, the isolated dimer alone does not display any assembly activity, agreeing with the previous observation that fresh monomers have to be added into the packaging intermediate in order to re-initiate the packaging process (Zhao et al., 2016; Zhao et al., 2017; Shu & Guo, 2003). Most hexameric AAA+ structures show a typical domain arrangement in which the nucleotide-binding pocket lies in the interface between two protomers. Such a structural arrangement supports the conclusion that in the active complex ring, the arginine finger of one subunit comes into close proximity to the nucleotide bound in the neighboring subunit. A hexameric ring of phi29 gp16 ATPase has also been modeled and aligned with that of the hexameric FtsK DNA translocase of Escherichia coli. The position of the arginine finger of one subunit of gp16 is shown to outstretch to the active site of a neighboring subunit, agreeing with the other ATPases that arginine finger was part of the ATP-binding pocket for cooperative behavior. With the above evidences, the cooperativity of subunit communication regulated by arginine finger has been illustrated. Oligomeric helicases contain one ASCE domain per monomer, with the ATP site at the interface between adjacent subunits, and rely on the interaction with neighboring subunits to provide the full nucleotide-binding pocket. In contrast, ATPases from SF1 and SF2 (Superfamily 1 and Superfamily 2) typically contain tandem ASCE folds and bind the nucleotide at the interface between the two domains, with the N-terminal providing the Walker A and Walker B motif, and the C-terminal providing other elements. The homologous type II transmembrane proteins LAP1 and LULL1 adopt nucleotide-free ATPase folds, and donate arginine fingers to complete the active sites of Torsin ATPases. The ring structure provides a central channel where the nucleic acid substrate is supposed to thread. A closer look at the GTP-binding site of the crystallized Ras/p120GAP/GDP AlF3 complex reveals that the closest distance between the fluoride groups of AlF3 and the amino group of Arg789 is around 2.6 Å, allowing for direct interactions between the arginine finger and the nucleotide in the transition state. The ATP-binding pocket of the SV40 LTag helicase (helicase superfamily III) is formed by two positively charged residues from the neighboring subunit, namely, Arg540 and Lys418. Interestingly, Lys418 rather than the conserved arginine finger residue Arg540 takes the location of that in the Ras/p120GAP pair (Wendler et al., 2012). As demonstrated above, arginine finger serves as a bridge between two independent subunits, thus forming a transient dimeric subunit. In wild-type gp16 ATPase, it was observed that both dimer and monomer forms were present in solution, as revealed by glycerol gradient centrifugation experiments (Zhao et al., 2016; Zhao et al., 2017). The communication between each two adjacent subunits mediated by the arginine finger results in an asymmetrical hexameric organization, which is supported by the asymmetrical structure in many other hexameric ATPase systems as shown by structural computation, X-ray diffraction, and Cryogenic electron microscopy (cryo-EM)imaging

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(Zhao et al., 2016; Arai et al., 2013; Stinson et al., 2015; Lyubimov et al., 2012; Ye et al., 2015). This phenomenon could provide some clues as to why the asymmetrical hexameric ATPase of gp16 of phi29 and gp17 of T4 was previously interpreted as having a pentameric configuration by Cryo-EM. Since the two adjacent subunits of the ATPase could interact with each other and form a closer dimer configuration, this dimer will appear as a monomeric subunit different from the other subunits and the hexameric ring as asymmetrical.

REFERENCES Arai, S., Saijo, S., Suzuki, K., Mizutani, K., Kakinuma, Y., Ishizuka-Katsura, Y., Ohsawa, N., Terada, T., Shirouzu, M., Yokoyama, S., Iwata, S., Yamato, I., Murata, T., 2013. Rotation mechanism of Enterococcus hirae V1-ATPase based on asymmetric crystal structures. Nature 493, 703–707. Chen, B., Sysoeva, T. A., Chowdhury, S., Guo, L., De, C. S., Hanson, J. A., Yang, H., Nixon, B. T., 2010. Engagement of arginine finger to ATP triggers large conformational changes in NtrC1 AAA+ ATPase for remodeling bacterial RNA polymerase. Structure 18, 1420–1430. Chen, C., Guo, P., 1997. Sequential action of six virus-encoded DNA-packaging RNAs during phage phi29 genomic DNA translocation. Journal of Virology 71, 3864–3871. De-Donatis, G., Zhao, Z., Wang, S., Huang, P. L., Schwartz, C., Tsodikov, V. O., Zhang, H., Haque, F., Guo, P., 2014. Finding of widespread viral and bacterial revolution dsDNA translocation motors distinct from rotation motors by channel chirality and size. Cell Biosci 4, 30. Elles, L. M., Uhlenbeck, O. C., 2008. Mutation of the arginine finger in the active site of Escherichia coli DbpA abolishes ATPase and helicase activity and confers a dominant slow growth phenotype. Nucleic Acids Research 36, 41–50. Hanson, P. I., Whiteheart, S. W., 2005. AAA+ proteins: have engine, will work. Nature Reviews Molecular Cell Biology 6, 519–529. Kolomeisky, A. B., Fisher, M. E., 2007. Molecular motors: a theorist’s perspective. Annual Review of Physical Chemistry 58, 675–695. Lyubimov, A. Y., Costa, A., Bleichert, F., Botchan, M. R., Berger, J. M., 2012. ATP-dependent conformational dynamics underlie the functional asymmetry of the replicative helicase from a minimalist eukaryote. Proceedings of the National Academy of Sciences of the United States of America 109, 11999–12004. 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, 446–450. Schwartz, C., De Donatis, G. M., Zhang, H., Fang, H., Guo, P., 2013. Revolution rather than rotation of AAA+ hexameric phi29 nanomotor for viral dsDNA packaging without coiling. Virology 443, 28–39. Shu, D., Guo, P., 2003. Only one pRNA hexamer but multiple copies of the DNA-packaging protein gp16 are needed for the motor to package bacterial virus phi29 genomic DNA. Virology 309(1), 108–113. Stinson, B. M., Baytshtok, V., Schmitz, K. R., Baker, T. A., Sauer, R. T., 2015. Subunit asymmetry and roles of conformational switching in the hexameric AAA+ ring of ClpX. Nature Structural & Molecular Biology 22, 411–416. Wendler, P., Ciniawsky, S., Kock, M., Kube, S., 2012. Structure and function of the AAA+ nucleotide binding pocket. Biochimica et Biophysica Acta 1823, 2–14. Ye, Q., Rosenberg, S. C., Moeller, A., Speir, J. A., Su, T. Y., Corbett, K. D., 2015. TRIP13 is a protein-remodeling AAA+ ATPase that catalyzes MAD2 conformation switching. Elife. 4, e07367. Zhao, Z., De-Donatis, G. M., Schwartz, C., Fang, H., Li, J., Guo, P., 2016. Arginine finger regulates sequential action of asymmetrical hexameric ATPase in dsDNA translocation motor. Molecular and Cellular Biology 36, 2514–2523. Zhao, Z., Zhang, H., Shu, D., Montemagno, C., Ding, B., Li, J., Guo, P., 2017. Construction of asymmetrical hexameric biomimetic motors with continuous single-directional motion by sequential coordination. Small 13, 1601600.

9

Mechanism of Revolving Motors Peixuan Guo and Zhengyi Zhao University of Kentucky

CONTENTS 9.1 Revolving Motion in Biological Motors ............................................................................... 103 9.2 One-Way Traffic of Revolving Biomotors ............................................................................ 106 References ...................................................................................................................................... 107

9.1

REVOLVING MOTION IN BIOLOGICAL MOTORS

ATPase is an enzyme that facilitates the hydrolysis of ATP, and gp16 in the phi29 dsDNA packaging motor converts the energy that results from this process into physical motion. The valve of the connector channel between the motor and procapsid is structured in such a way that it allows the dsDNA to enter the chamber but not exit. A virus-encoded packaging RNA, or pRNA, is required for the translocation of dsDNA into the phi29 procapsid. It coordinates the action of the separate components of gp16 and the central channel, making for an integrated motor. When ATP interacts with gp16, a conformational change in gp16 increases its binding to dsDNA. Finally, the hydrolysis of ATP functions to force dsDNA through the channel. The FtsK motor domain from Pseudomonas aeruginosa is a sixfold symmetric ring, with ADP bound in every subunit (Massey et al., 2006) as revealed by crystallization studies. The same study also presented a monomeric crystal form of the motor domain with ATP bound. When the b domains of these two structures were superimposed, their a domains were shifted relative to each other in a hinge-like opening, which was able to move the point in the a domain juxtaposed to the DNA substrate by 5.5 Å (equivalent to 1.6 bp). The translocation of 1.6 bp of dsDNA per subunit of FtsK (Massey et al., 2006; Crozat & Grainge, 2010) might be correlated with ATP hydrolysis. It strongly agrees with the step size of the phi29 DNA packaging motor, in which one ATPase subunit uses one ATP each time to package 1.75 nucleotides (Guo et al., 1987; Schwartz et al., 2013b; Schwartz et al., 2013a; Zhao et al., 2013; Guo et al., 2013), and also that in bacteriophage T3 with 1.8 bp per ATP (Morita et al., 1993). Based on that, it has been proposed that FtsK possesses a rotary inchworm mechanism (Massey et al., 2006; Crozat & Grainge, 2010), with ATPase subunits acting sequentially. In the model proposed by Massey et al., each hydrolyzed ATP moves the DNA by 2 bp (~6.8 Å, larger than the 5.5 Å difference between the two crystal forms as discussed above) in an inchworm-like movement with the central DNA having contact with both end domains. The relative strength of interaction between each domain and the DNA is dependent upon the ATP binding or hydrolysis state; upon ATP hydrolysis, one DNA contact is lost, while the other contact is strong but shifted by ~6.8 Å, resulting in net movement of 2 bp of DNA along the central channel (Guo et al., 2016). An integer number of bases was chosen as the most likely model as it allows each monomer in the ring to contact the dsDNA in the same manner at every subunit, whether this is with the repeating sugar-phosphate backbone or the bases themselves. Movement by a non-integer number of bases means that the protein-DNA contact necessary for movement of the substrate would be different in any two adjacent subunits (Guo et al., 2016). The movement of dsDNA by one subunit also functions to bring the next monomer DOI: 10.1201/9780429203367-10

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in the ring into contact with the DNA, handing the DNA substrate on to the adjacent monomer with minimal rotation. The next subunit also translocates 2 bp of DNA, and so on around the ring. The result is that 12 bp of DNA is translocated per catalytic cycle of the hexamer where all six monomers hydrolyze ATP once. This figure is close to the 10.5 bp per helical turn in B-form DNA, with the extra 1.5 bp appearing as a slight rotation. If the protein is anchored (both FtsK and SpoIIIE and membrane bound at their N-termini), this results in the generation of positive DNA supercoiling ahead of the motor and negative behind. Indeed, induction of supercoiling has been seen in bulk biochemical assays (Aussel et al., 2002) and in single-molecule experiments (Saleh et al., 2005). The observed supercoiling induction for FtsK translocation of one +ve supercoil ahead of the motor per 150 bp translocated is in broad agreement with the rotary inchworm model (Guo et al., 2016). The rotary inchworm model adopted for the FtsK motor also proposed an obligatory hand-off event between adjacent monomers within a single ring, such that the presence of a single catalytically inactive subunit would effectively inactivate the entire hexamer (Guo et al., 2016). This was backed up by biochemical data: mutants that were unable to bind ATP were mixed in different ratios with wild-type subunits, and the relative ATPase activity was compared to wild type only. With an increasing amount of mutant subunits, ATPase activity decreased rapidly, confirming the predicted pattern (Guo et al., 2016). Interestingly, a fusion protein has been produced in which three linked motor domains, effectively a covalent trimer of FtsK, were contained in a single polypeptide. This construct was found to be a very active DNA translocase motor (Guo et al., 2016). Within it, the Walker A and Walker B motifs, for nucleotide binding and hydrolysis, respectively, could be mutated in specific subunits, leading to the surprising finding that a single active-site mutant, or two non-adjacent mutants per hexamer, did not cause a great decrease in ATPase activity or the speed of translocation along dsDNA. However, the presence of these mutations did reduce the ability of the hexamer to produce force as judged by the ability of the protein to displace either protein or DNA triplexes (Guo et al., 2016). It is important to consider that when the ATPase subunits were fused into a concatemer, the ATPase could refold or form a higher-order multimer, rendering the interpretation of the results challenging. In addition, the linking of subunits as a concatemer may relax the otherwise strict inter-subunit interactions that coordinate the binding and hydrolysis of ATP in the protein (such as the role of the arginine finger that stretches between one monomer and the active site of its neighbor). In order to explain these results, a new model was proposed in which more than one subunit within a hexameric ring would contact DNA concurrently, based upon the escort or “spiral staircase” model of Rho and E1 helicases (Guo et al., 2016). In these hexameric helicases, multiple subunits can contact the DNA/RNA substrate at the same time, with the single contact point for each monomer being at a different level around the ring, rather like the stairs in a spiral staircase. ATP hydrolysis forcefully moves one of the contact points downwards through the ring, and the other contacts move along passively. When the last contact point at the bottom of the ring is reached, the protein arm becomes free and can then move back up the top to re-engage with the polynucleotide substrate and reinitiate the cycle of movement down the staircase (Guo et al., 2016). With a flexible and compressible single-stranded DNA/RNA substrate, the movement of the protein-DNA contacts is small enough that the protein can remain with the DNA/RNA through a full catalytic cycle of every subunit in the ring. However, dsDNA is a much stiffer and non-compressible substrate than ssDNA or RNA, so a DNA translocase would have immense trouble utilizing an identical mechanism. Within a DNA translocase channel, each single protein contact would have to move almost 30 Å to maintain contact with the DNA during a full catalytic cycle around the hexameric ring. This amount of movement seems unlikely (Guo et al., 2016). However, in the channel of the DNA packaging motor of all dsDNA bacteriophages, a 30° left-handed channel structure was found. Such a left-handed configuration might assist to compensate for such constraints during shifting of the dsDNA and maintain the balanced contact during the advance of the dsDNA. Furthermore, it would

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be energetically unfeasible to compress the double helix greatly yet still produce power from ATP hydrolysis. Therefore, an intermediate or “limited escort” model was proposed for FtsK whereby three adjacent monomers in a ring contact dsDNA simultaneously. This could allow for one inactive subunit to be skipped if the monomers on either side of it are able to translocate. It would also imply that two adjacent mutant subunits would produce an inactive hexamer, which is consistent with the data (Guo et al., 2016). Recent results for SpoIIIE also support an escort model (Liu et al., 2015); using a single-molecule setup with an optical trap, it was found that SpoIIIE most likely has a step size on DNA of 2 bp, and that (at least) two adjacent subunits must be able to contact DNA simultaneously (Liu et al., 2015). It is noteworthy that this model is based on hexameric SpoIIIE made of six monomers, not linked multimers as with the FtsK experiments above, removing the concern of how the linked multimers may form an active translocase. Together, these studies suggest a highly coordinated cycle of ATP binding and hydrolysis around the ring, with monomers becoming active and hydrolyzing ATP sequentially. Yet the hexameric rings retain the ability to bypass individual inactive monomers by virtue of having multiple concurrent DNA contacts (two or three monomers contacting DNA at the same time). Both the inchworm and partial escort models are largely consistent with a revolving motor mechanism. They propose that dsDNA touches the internal surface of the hexameric ring and that the contact point between protein and DNA revolves around the inner surface of the protein multimer with minimal rotation (Guo et al., 2016). They also suggest that a defined number of bases would be moved per hydrolyzed ATP; if each subunit contacts the DNA substrate in an identical fashion around the ring, then the DNA must be moved by a defined length at each step to maintain constant interaction. A slight twisting of the DNA at each step is then necessary to maintain the contacts with the dsDNA around the ring, preserving the register between DNA and protein. While the angle between adjacent subunit active sites in a hexamer is 360°/6 or 60°, the angle between adjacent phosphates around the dsDNA axis is 360°/10.5 (about 34°) (Guo et al., 2016). If precisely 2 bp are translocated per subunit, the DNA must twist an extra 8o to maintain the identity of each protein-DNA contact. DNA within cells is negatively supercoiled, with a supercoiling density in Escherichia coli of around −0.05. Thus, the amount of twisting required for each FtsK power step is reduced to around 5° per 2 bp translocated, which corresponds to one supercoil induced for every 144 bp translocated (Guo et al., 2016). This theoretical value is almost identical to the value of one supercoil per 150 bp observed using single-molecule experiments. If this is the case, small over-winding of the DNA ahead of a translocating FtsK will produce positive supercoiling ahead of the protein; however, if the left-handed configuration as observed in the DNA packaging motor of dsDNA bacteriophages exists, then the supercoiling will not have occurred. On the other hand, in the cell this might be removed by either the action of DNA gyrase or the occasional slipping of the motor to release the torsional tension in the DNA (Guo et al., 2016). Furthermore, SpoIIIE is hypothesized to gain directionality through the recognition of skewed 8-bp sequences in an analogous manner to FtsK. The SpoIIIE recognition sequence (SRS) of Bacillus subtilis is also a skewed 8-bp sequence (GAGAAGGG) and is bound by the C-terminal domain. However, recent models propose a subtly different mechanism for ensuring that SpoIIIE translocates DNA in the desired direction: rather than SRS acting as a preferential loading site onto the DNA, as proposed for FtsK (Grainge et al., 2011). SpoIIIE may randomly associate with DNA and scan for SRSs without the motor ATPase being active by a sliding/hopping mechanism. It has also been suggested that the γ domain of SpoIIIE acts to inhibit motor activity (Besprozvannaya et al., 2013) and that subsequent encounter of the passively sliding/hopping motor with an SRS in the correct orientation relieves this repression. An alternative hypothesis is that the encounter of SRS leads to a conformational change in SpoIIIE, converting the inactive “open hexamer,” which is capable of rapidly dissociating from DNA, to a stable closed form, in which ATPase activity is now greatly activated (Cattoni et al., 2013; Cattoni et al., 2014). Both FtsK and SpoIIIE can bind

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to random DNA, and ATPase activity is not dependent upon the presence of KOPS/SRS. However, the interaction of the motor protein and its cognate DNA sequence leads to motor activation, either by loading a hexamer, by stabilizing a closed form of the hexamer on the DNA, by relieving motor inhibition, or perhaps by a combination of these mechanisms. Whether FtsK and SpoIIIE in fact respond to their respective directionality sequences differently or whether the two related proteins behave in a similar fashion remains to be seen.

9.2

ONE-WAY TRAFFIC OF REVOLVING BIOMOTORS

As reported, revolving biomotors drive their substrate translocation in a one-way traffic manner under the coordination of the motor components. There are several factors that contribute to the directional translocation: the loop and the four lysine rings aligned within the connector inner wall, ATP-coupled conformational entropy alternations, left-handed configuration of the connector subunit, and the recognition of 5′-3′ strand for the single-directional movement. The one-way flow loops within the connector channel of the phi29 dsDNA packaging motor is one of the most important factors determining the advancement of genome towards the capsid without going back (Fang et al., 2012; Zhao et al., 2013; Geng et al., 2013). Both biochemical and biophysical experiments with loop-deleted connectors have showed the ability of two-way traffic of the DNA substrate (Fang et al., 2012; Geng et al., 2011; Grimes et al., 2011; Serwer, 2010; Isidro et al., 2004), proving the role of these inner loops as a ratchet or clamp during DNA translocation, in line with the “push-through-one-way-valve” model (Fang et al., 2012; Zhao et al., 2013; Zhang et al., 2012; Jing et al., 2010). Structural studies of the SPP1 bacteriophage connector channel have shown the close proximity of its inner loops to the dsDNA substrate via non-ionic interactions (Orlova et al., 2003) and the high similarity between SPP1 and phi29 protein channels (Chai et al., 1994), indicating the conserved role of the loops during genome packaging. Besides, the conserved 30° left-handed twist of the channel, which is anti-chiral to the right-handed dsDNA substrate, will compensate for the angle and distance difference by each contact between dsDNA and the channel, supporting the one-way revolving motion without rotation. During packaging, the dsDNA genome is processed by contact of the connector with one strand of DNA in the 5′ to 3′ direction, as modification of only that strand abolished dsDNA packaging (Aathavan et al., 2009; Oram et al., 2008). Extensions up to 12 bases at the end is tolerable, while extensions to 20 or more bases significantly blocked the DNA packaging of the T4 motor, agreeing with the previous conclusion since one complete circle of revolving is equal to one helical turn of 10.5 base pairs of dsDNA. In addition, force generation from ATPase also contributes to the oneway movement. As described in the early section, ATP binding and hydrolysis induce two consecutive entropic and conformational alternations of ATPase with either a high or low affinity for DNA, with the arginine finger motif as a mediator for signal transfer, which results in a sequential action facilitating the translocation of the DNA substrate in one direction. Four lysine residues are reported to be aligned within each subunit of the phi29 connector interior surface (Guasch et al., 2002). Similar patterns have also been observed in the phages of SPP1, P22, and phi29 (Guasch et al., 2002; Simpson et al., 2000). These electropositive lysines form four relaying rings and interact with the electronegative DNA phosphate backbone, generating an auxiliary force involved in genome packaging (Fang et al., 2012; Schwartz et al., 2013a; Schwartz et al., 2013b; Zhao et al., 2013). This resulting force leads to uneven speed alternations during the DNA translocation, mostly with four pauses (Chistol et al., 2012; Schwartz et al., 2013b; Zhao et al., 2013), as previously reported in both phi29 (Chistol et al., 2012; Moffitt et al., 2009; Moffitt et al., 2009) and T4 (Kottadiel et al., 2012; Kottadiel et al., 2012). The effects of the lysine layers on genome translocation can be interpreted using phi29 biomotor as an example (Zhao et al., 2013). Vertically, these four lysine layers of phi29 fall within 3.7 nm (Guasch et al., 2002) inside the 7-nm connector channel, which is an average of ~0.9 nm apart between each two rings. During the revolution of DNA inside the motor channel, the

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distance variation due to the mismatch between genome base (10.5 bp for 360°) and the channel subunits (12 subunits) can be compensated by interactions between dsDNA phosphate backbone and the positively charged lysine from the next subunit, leading to a slight pause in DNA advancement. Continuation of the interactions leads to the four pauses during packaging as mentioned above.

REFERENCES Aathavan, K., Politzer, A. T., Kaplan, A., Moffitt, J. R., Chemla, Y. R., Grimes, S., Jardine, P. J., Anderson, D. L., Bustamante, C., 2009. Substrate interactions and promiscuity in a viral DNA packaging motor. Nature 461, 669–673. Aussel, L., Barre, F-X., Aroyo, M., Stasiak, A., Stasiak, A. Z., Sherratt, D., 2002. Cell 108,195–205. Besprozvannaya, M., Pivorunas, V. L., Feldman, Z., Burton, B. M., 2013. Journal of Biological Chemistry 4; 288(40), 28962–28974. Cattoni, D.I., Chara, O., Godefroy, C., Margeat, E., Trigueros, S., Milhiet, P.E., Nöllmann, M., 2013. EMBO Report 14, 473–479. Cattoni, D. I., Thakur, S., Godefroy, C., Le Gall, A., Lai-Kee-Him, J., Milhiet, P. E., Bron, P., Nöllmann, M., 2014. Nucleic Acids Research 42(4), 2624–2636. Chai, S., Kruft, V., Alonso, J. C., 1994. Analysis of the Bacillus subtilis bacteriophages SPP1 and SF6 gene 1 product: a protein involved in the initiation of headful packaging. Virology 202, 930–939. Chistol, G., Liu, S., Hetherington, C. L., Moffitt, J. R., Grimes, S., Jardine, P. J., Bustamante, C., 2012. High degree of coordination and division of labor among subunits in a homomeric ring ATPase. Cell 151, 1017–1028. Crozat, E., Grainge, I., 2010. FtsK DNA translocase: the fast motor that knows where it’s going. Chembiochem 11, 2232–2243. Fang, H., Jing, P., Haque, F., Guo, P., 2012. Role of channel lysines and “push through a one-way valve” mechanism of viral DNA packaging motor. Biophysical Journal 102, 127–135. Geng, J., Wang, S., Fang, H., Guo, P., 2013. Channel size conversion of phi29 DNA-packaging nanomotor for discrimination of single- and double-stranded nucleic acids. ACS Nano 7, 3315–3323. Geng, J., Fang, H., Haque, F., Zhang, L., Guo, P., 2011. Three reversible and controllable discrete steps of channel gating of a viral DNA packaging motor. Biomaterials 32, 8234–8242. Grainge, I., Lesterlin, C., Sherratt, J. D., 2011, Nucleic Acids Research 39, 5140–5148. Grimes, S., Ma, S., Gao, J., Atz, R., Jardine, P. J., 2011. Role of phi29 connector channel loops in late-stage DNA packaging. Journal of Molecular Biology 410, 50–59. Guasch, A., Pous, J., Ibarra, B., Gomis-Ruth, F. X., Valpuesta, J. M., Sousa, N., Carrascosa, J. L., Coll, M., 2002. Detailed architecture of a DNA translocating machine: the high-resolution structure of the bacteriophage phi29 connector particle. Journal of Molecular Biology 315, 663–676. Guo, P., Noji, H., Yengo, C. M., Zhao, Z., Grainge, I., 2016. Biological nanomotors with revolution, linear, or rotation motion mechanism. Microbiology and Molecular Biology Reviews 80, 161–186. Guo, P., Peterson, C., Anderson, D., 1987. Prohead and DNA-gp3-dependent ATPase activity of the DNA packaging protein gp16 of bacteriophage phi29. Journal of Molecular Biology 197, 229–236. Guo, P., Schwartz, C., Haak, J., Zhao, Z., 2013. Discovery of a new motion mechanism of biomotors similar to the earth revolving around the sun without rotation. Virology 446, 133–143. Isidro, A., Henriques, A. O., Tavares, P., 2004. The portal protein plays essential roles at different steps of the SPP1 DNA packaging process. Virology 322, 253–263. Jing, P., Haque, F., Shu, D., Montemagno, C., Guo, P., 2010. One-way traffic of a viral motor channel for double-stranded DNA translocation. Nano Letters 10, 3620–3627. Kottadiel, V. I., Rao, V. B., Chemla, Y. R., 2012. The dynamic pause-unpackaging state, an off-translocation recovery state of a DNA packaging motor from bacteriophage T4. Proceedings of the National Academy of Sciences of the United States of America 109, 20000–20005. Liu, N., Chistol, G., Bustamante, C., 2015. eLife 4, e09224. Massey, T. H., Mercogliano, C. P., Yates, J., Sherratt, D. J., Lowe, J., 2006. Double-stranded DNA translocation: structure and mechanism of hexameric FtsK. Molecular Cell 23, 457–469. 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, 446–450. Morita, M., Tasaka, M., Fujisawa, H., 1993. DNA packaging ATPase of bacteriophage T3. Virology 193, 748–752.

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Oram, M., Sabanayagam, C., Black, L. W., 2008. Modulation of the packaging reaction of bacteriophage T4 terminase by DNA structure. Journal of Molecular Biology 381, 61–72. Orlova, E. V., Gowen, B., Droge, A., Stiege, A., Weise, F., Lurz, R., van, H. M., Tavares, P., 2003. Structure of a viral DNA gatekeeper at 10 Å resolution by cryo-electron microscopy. EMBO Journal 22, 1255–1262. Saleh, O.A., Bigot, S., Barre, F.X., Allemand, J.F., 2005. Nature Structural & Molecular Biology 12, 436–440. Schwartz, C., De Donatis, G. M., Fang, H., Guo, P., 2013a. The ATPase of the phi29 DNA-packaging motor is a member of the hexameric AAA+ superfamily. Virology 443, 20–27. Schwartz, C., De Donatis, G. M., Zhang, H., Fang, H., Guo, P., 2013b. Revolution rather than rotation of AAA+ hexameric phi29 nanomotor for viral dsDNA packaging without coiling. Virology 443, 28–39. Serwer, P., 2010. A hypothesis for bacteriophage DNA packaging motors. Viruses 2, 1821–1843. Simpson, A. A., Tao, Y., Leiman, P. G., Badasso, M. O., He, Y., Jardine, P. J., Olson, N. H., Morais, M. C., Grimes, S., Anderson, D. L., Baker, T. S., Rossmann, M. G., 2000. Structure of the bacteriophage phi29 DNA packaging motor. Nature 408, 745–750. Zhang, H., Schwartz, C., De Donatis, G. M., Guo, P., 2012. “Push through one-way valve” mechanism of viral DNA packaging. Advances in Virus Research 83, 415–465. Zhao, Z., Khisamutdinov, E., Schwartz, C., Guo, P., 2013. Mechanism of one-way traffic of hexameric phi29 DNA packaging motor with four electropositive relaying layers facilitating anti-parallel revolution. ACS Nano 7, 4082–4092.

10

Mechanism of Rotary Motors Peixuan Guo and Zhengyi Zhao University of Kentucky

CONTENTS 10.1 Rotation Motion in F1 ........................................................................................................... 109 10.1.1 Single-Molecule Rotation Assay of F1 ...................................................................... 109 10.1.2 Torque of F1............................................................................................................... 110 10.1.3 Chemomechanical Coupling of F1 ............................................................................ 110 10.1.4 Torque Generation Steps of F1 .................................................................................. 111 10.1.5 Critical Role of Phosphate-Binding Sites in Force Generation ................................ 112 10.2 Rotation Motion in Fo ........................................................................................................... 113 References ...................................................................................................................................... 114

10.1 10.1.1

ROTATION MOTION IN F1 single-MoleCule Rotation assay oF F1

In FoF1 ATPase synthase, ATP synthesis from ADP and inorganic phosphate (Pi) is the basic reaction for cell energy production in animals, plants, and microorganisms. Cells have two major ATP production pathways. One is substrate-level ATP synthesis reaction like glycolysis, and the second is oxidative phosphorylation where electron transfer among electron carriers down the redox potential and drives respiratory chain proteins to pump protons for proton motive force (pmf) generation. ATP is synthesized upon the backflow of protons down pmf. FoF1 ATP synthase catalyzes this reaction as the terminal reaction of oxidative phosphorylation. Dynamic behavior of the ATP-driven rotation of F1 is well characterized in single-molecule rotation assay where the α3β3 stator ring is immobilized on a coverslip and the rotation probe is attached onto the outwardly protruding part of the γ subunit of F1 (Guo et al., 2016). In the presence of ATP, F1 rotates the γ subunit counterclockwise as viewed from the Fo side. Since the unidirectional rotation of F1 was first visualized from Bacillus PS3, several kinds of rotary ATPases – F1’s from Escherichia coli (Hisabori et al., 1999), chloroplast from spinach (Hisabori et al., 1999), human mitochondria (T. Suzuki, personal communication), and V-ATPases from Thermus thermophiles (Imamura et al., 2003) and Enterococcus hirae (Minagawa et al., 2013) – were examined in the rotation assay. All rotary ATPases show the counterclockwise rotation without exception, implying a highly conserved structural basis for unidirectional rotation. Among them, F1 from Bacillus PS3, termed TF1 (thermophilic F1), is the best characterized in terms of rotary dynamics. Therefore, the rotation features of F1 introduced hereafter are based on the findings in the single-molecule rotation assay of TF1 unless mentioned. Rotation of the isolated F1 motor driven by ATP hydrolysis was directly observed with an optical microscope (Guo et al., 2016; Okuno et al., 2011; Noji et al., 1997). Consistent with the pseudo-3fold symmetry of the α3β3 stator ring, F1 rotates γ in discrete 120° steps (Yasuda et al., 1998), each coupled with a single turnover of ATP hydrolysis (Yasuda et al., 1998; Rondelez et al., 2005). An intermediate state was also observed after ATP binding and before hydrolysis at an 80° angle from the ATP-waiting angle (Guo et al., 2016; Shimabukuro et al., 2003). Therefore, each 120° step can be resolved into 80° and 40° substeps. Recent reaction scheme suggests that the 80° substep is driven by ATP binding and ADP release that occur on different β subunits, and that the 40° substep DOI: 10.1201/9780429203367-11

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is initiated after hydrolysis of bound ATP and Pi release that also occur on different β subunits (Guo et al., 2016; Watanabe et al., 2010). It is under debate as to which reaction state the crystal structure of F1 represents; although several lines of experiments suggest that most crystal structures of F1 represent the intermediate state before the 40° substep (hydrolysis-waiting state), there are some crystal structures that are not explainable on this basis (Guo et al., 2016; Menz et al., 2001; Stracke et al., 2000).

10.1.2

toRque oF F1

Rotary torque that F1 generates is estimated from the viscous friction against the rotating probe: drag coefficient times angular velocity gives the torque that F1 generates. Although some groups reported higher torque for F1 from E. coli (EF1) (Spetzler et al., 2006; Rees et al., 2012), most data show that F1 generates a torque of 40 pN nm irrespective of species (Yasuda et al., 1998; Panke et al., 2001; Noji et al., 1999). A theory of non-equilibrium physics, fluctuation theorem, was also employed as a new analytical method to estimate torque generation. This theorem also showed that F1 generates a torque of 40 pNnm (Hayashi et al., 2012). Considering that the torque is generated at the stator-rotor interface 1 nm from the rotation axis, the generated force is 40 pN, much larger than conventional linear motors like myosin and kinesin (Kinosita, Jr. et al., 1998). Interestingly, V-ATPase is known to generate slightly smaller torque than F1, around 35 pN (Imamura et al., 2003). The magnitude of torque might reflect some physiological difference. Torque times angular displacement gives the work done upon rotation. Therefore, the work done by F1 per ATP is estimated to be 80 pNnm (40 pNnm × 2π/3) (Yasuda et al., 1998). This value roughly agrees with the released free energy upon single turnover of ATP hydrolysis under physiological conditions, consistent with highly efficient energy conversion of FoF1 ATP synthase. Another important finding about torque generation is that F1 exerts a constant torque of 40 pNnm irrespective of viscous drag coefficient (Yasuda et al., 2001). This means that even when a large viscous marker significantly extends the time scale of mechanical rotation by 1000 times or more, F1 bears high viscous drag force against a rotating probe, exerting constant torque. The rotation against large viscous drag is very similar to the situation under physiological conditions where the rotation torque of F1 and Fo is mostly balanced, although the situation is not exactly the same from a thermodynamic point of view. If F1 does not bear stall force condition and consumes ATP without rotation, FoF1 ATP synthase cannot interconvert pmf and free energy of ATP with high efficiency. Thus, robust torque generation against the high viscous friction is also relevant to the highly efficient energy conversion mechanism.

10.1.3

CHeMoMeCHaniCal Coupling oF F1

To achieve high energy conversion efficiency, high reversibility of chemomechanical coupling is required in addition to robust torque generation. The reversely coupled reaction of F1, ATP production upon clockwise rotation was tested by forcibly rotating F1 in a clockwise direction with magnetic tweezers. In the first experiment, a large number of F1 molecules immobilized on a coverslip were simultaneously rotated and the synthesized ATP was detected with chemiluminescent reaction (Itoh et al., 2004). However, because the exact number of active F1 molecules was not clear, the coupling efficiency was not determined. In the following experiment, actively rotating F1 molecules were individually encapsulated in a femtoliter chamber and rotated with magnetic tweezers (Rondelez et al., 2005). Due to the ultra-small reaction volume, a small number of synthesized ATP molecules significantly increased ATP concentration in the chamber. After being released from forcible rotation, F1 molecules resumed active ATP-hydrolyzing rotation that allowed for ATP measurement because the rotation velocity of F1 is proportional to ATP concentration under the condition of low ATP concentration. This experiment showed that F1 synthesizes ATP with high efficiency in the presence of the ε subunit. Although the role of the ε subunit has been extensively

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studied, it still remains elusive as to how the ε subunit enhances the coupling efficiency of ATP synthesis reaction of F1. As mentioned above, the step size of F1 rotation is 120°, each of which is driven by a single turnover of ATP hydrolysis. However, it does necessarily mean that each 120° step is driven by a particular β subunit. The actual reaction scheme of F1 is much more complex. While all β subunits participate in torque generation for each 120° step, each β subunit is in a distinct conformational and thereby catalytic state (Abrahams et al., 1994). In addition, the 120° step is further resolved into 80° and 40° substeps, as shown by kinetic analysis. The 80° substep is initiated after ATP binding and ADP release. The 40° substep is initiated after hydrolysis of bound ATP and Pi release. The dwell angles before the 80° and 40° substeps are referred to as ATP-waiting angle and hydrolysis angle, respectively. To solve the puzzling reaction scheme, elaborated single-molecule experiments were conducted, such as simultaneous observation of fluorescently labeled ATP and rotation (Adachi et al., 2007), rotation assays with hybrid F1 from the wild type and mutant β subunits (Ariga et al., 2007), and single-molecule manipulation experiments (Watanabe et al., 2010). Most parts of the reaction scheme have been solved (Okuno et al., 2011). The recent reaction scheme is as follows: each β binds to ATP when the γ subunit is oriented to a particular ATP-waiting angle (defined as 0°). Then, β hydrolyzes bound ATP when the γ subunit rotates 200°. ADP inorganic phosphates (Pi) are released at 240° and at 320°. The other 2 β subunits also undergo the same reaction pathway, while the phase of the catalytic state varies by +120° or −120°. Therefore, each 80° substep is initiated after ATP binding and ADP release that occur at different β subunits, and the 40° substep is initiated after hydrolysis and Pi release that also occur at different β subunits. It should be noted that the exact angle for Pi release is still under debate (Shimo-Kon et al., 2010; Stracke et al., 2000; Okazaki & Hummer, 2013). Many experiments have showed that the large conformational change occurred in the catalytic β subunit (Abrahams et al., 1994; Kobayashi et al., 2010; Masaike et al., 2008). Upon nucleotide binding, β subunit undergoes the opened-to-closed conformational transition (Uchihashi et al., 2011), thus pushing the γ subunit as shown by nuclear magnetic resonance (NMR) study (Kobayashi et al., 2010), single fluorophore imaging (Masaike et al., 2008), and atomic force microscopy (AFM) data, as expected from the crystal structure of MF1. The elementary catalytic steps of ATP binding, hydrolysis, and release of ADP and Pi are believed to serve as a major torque-generating step.

10.1.4

toRque geneRation steps oF F1

Large conformational transition does not necessarily guarantee large force generation. To estimate the contribution of ATP-binding and hydrolysis steps for torque generation in F1, single-molecule stall experiments were conducted where the equilibrium constants of ATP binding and hydrolysis were determined as a function of the rotary angle (Watanabe et al., 2012). As a result, it was revealed that the equilibrium constant of ATP binding exponentially increases with the forward rotation of the γ subunit. This means that the F1-ATP complex is progressively stabilized upon the rotation of γ, progressively releasing binding energy via γ rotation. This affinity change process is a typical induced-fit. Although the hydrolysis process also showed the angle-dependent stabilization of the post-hydrolysis state with the rotation of γ, the magnitude of equilibrium change was significantly small compared to the ATP-binding step. The estimated contribution of ATP binding for torque generation during the 80° substep is around 50% (up to 67%). The remaining torque generation is attributed to ADP release from a different β subunit. The contribution of hydrolysis in torque generation during the 40° substep is at most 21%, and Pi release is thought to be the major torque generation step in the 40° substep. Thus, it is evident that the role of chemical cleavage of phosphoester bond on the catalytic site in force generation is minor in F1. The stall experiment also showed that the conformational transition upon the induced-fit process after ATP binding is one of the major torque generation steps. It is highly likely that the induced-fit accompanies the openedto-closed transition predicted from the crystal structure of MF1. The torque generation upon ADP

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release would be induced-unfit, triggering closed-to-open transition. The conformational change upon Pi release is unclear. One feasible scenario is that the opening of the interface between the α and β subunits is coupled with Pi release (Ito et al., 2013). As mentioned above, the induced-fit process following the first ATP docking to the catalytic site on the β subunit is a major torque-generating process of F1. According to the crystal structure of MF1, adenine ring is surrounded by aromatic amino acids, while the phosphate moiety of ATP forms many hydrogen bonds with surrounding residues. Although crystal structures provide atomic pictures of conformations before and after conformational transition, it remained elusive as to which interactions are responsible in triggering the induced-fit among adenine ring, ribose, and phosphoester of ATP. Recently, the role of adenine ring in torque generation was tested in rotation assay using a synthetic, base-free nucleotide (Arai et al., 2014). The impact of depletion of adenine ring was remarkable in the kinetics of rotation; the binding rate constant was decreased by 6 orders of magnitude. However, the synthetic base-free nucleotide supports powerful rotation of F1 similar to ATP; generated torque was not different between ATP-driven rotation and base-free nucleotidedriven rotation. Thus, it was proven that the interaction between adenine ring and the catalytic site is not responsible for torque generation. The principal role of adenine ring is rate enhancement of the first docking process, but it is not involved in the induced-fit process.

10.1.5

CRitiCal Role oF pHospHate-binding sites in FoRCe geneRation

The role of phosphoester in force generation was examined by the use of mutant F1’s, in which mutation is introduced at the phosphate-interacting residues. Targeted residues are P-loop lysine, glutamic acid of general base, and arginine finger, all of which are well known to be critical for retaining the catalytic power of F1. Mutations at these residues lower the catalytic activity of F1 down to an undetectable level in biochemical assay. P-loop lysine, the lysine in the highly conserved phosphate-binding loop (P-loop with the common sequence GXXXXGKT/S) among NTPases, is well known to be the catalytically most critical residue in the P-loop; it is the common target to knock out ATPase activity of many ATPases. Glutamic acid of general base is also well conserved among ATPases that interact with the γ phosphate via a water molecule. Glutamic acid of general base has been thought to activate the intervening water molecule to induce nucleophilic attack of the water molecule to the γ phosphate (Guo et al., 2016). Recent quantum chemical calculation and single-molecule analysis revealed that the role of general base is not water activation but the enhancement of proton transfer from phosphoester to bulk solution (Hayashi et al., 2012). Arginine finger is also a highly conserved arginine residue among G proteins, AAA proteins, and RecA-type ATPases including F1. Arginine finger is located at an interface of nucleotide-binding subunits (Guo et al., 2016). ATP binds the interface of two subunits, one of which possesses the majority of ATPbinding residues, while the other has little but catalytically critical residue that is arginine finger (Guo et al., 2016). The best-characterized arginine finger is that of G-protein-activating protein (GAP). F1 has arginine finger on the α subunit. The crystal structure of MF1 with chemical analog of the γ phosphate suggests that arginine finger stabilizes the transient state of hydrolysis reaction, which was supported by biochemical (Nadanaciva et al., 1999), theoretical (Dittrich et al., 2003), and single-molecule studies (Komoriya et al., 2012). The role of P-loop lysine, glutamic acid of general base, and arginine finger in torque generation was examined in the rotation assay with alanine mutants, in which alanine substitution was introduced at one of the target residues (Watanabe et al., 2014). Although all mutant F1’s support continuous and unidirectional rotation, the mutants showed significantly slower rotation than the wild type; the slowest was alanine mutant at P-loop lysine (1/1,000 of the wild type). Thus, the role of mutations at phosphate-binding residues on kinetic power is significant as well as that of adenine ring. Prominent impact of the mutations was found in torque generation; all mutants were deficient for efficient torque generation. Alanine substitution at the general base and arginine finger halved torque. Mutations of P-loop lysine decreased torque down to 25% of the wild type.

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Thus, phosphate-binding sites play a critical role in rate enhancement of both catalysis and torque generation. Taking into account that the induced-fit process is a major torque-generating step, it was proposed that the progressive hydrogen bond formation between the phosphate moiety of bound ATP and the catalytic residues is the main driving force of the induced-fit-type conformational change: the opened-to-closed transition of the β subunit. The accompanying swing motion of the C-terminal domain of the β subunit would push the γ subunit to induce the rotation.

10.2

ROTATION MOTION IN FO

Two-hemi-channel model of Fo: The rotation of the c-ring against the a1b2 complex of Fo is driven by proton flow. Some bacteria have Fo fueled by sodium motive force (Meier et al., 2005). As the whole structure of Fo is not known, only a part of the ion-conducting pathway is revealed. Biochemical work has identified several charged residues in the transmembrane helices of the a and c subunits that would be directly involved in proton translocation. Among them, Asp or Glu of the c subunit resides in the middle of the C-terminal helix, and Arg of the a subunit, corresponding to cAsp61 and aArg210 of E. coli Fo, are highly conserved among species and thought to play crucial roles in proton translocation. The crystal structure of the c-ring from Ilyobacter tartaricus Fo, a Na+-transporting Fo, showed that the critical cGlu65 residues are occupied by Na+ ions (Meier et al., 2005). This finding established that this conserved carboxyl residue is one of the ion-binding sites. However, other charged residues are not found in the c subunit in the vicinity of the carboxyl residue, suggesting that the a subunit has other parts of the ion-conducting pathway. The most widely accepted model on proton translocation in Fo is the two-channel model, originally proposed by Junge et al. (1997). This model assumes that the a subunit possesses 2 hemi-channels, each of which spans half of the membrane but toward different sides. The hemi-channels connect the ionconducting carboxyl residue of the c subunit. Each channel is in contact with different c subunits, which are adjacent to each other. Thus, the a subunit interacts with 2 c subunits, each contacting via a different half channel of the a subunit. The proposed mechanism of proton transfer in the ATP synthesis mode is as follows: a proton enters the half channel exposed to the periplasmic side (or intermembrane space of the mitochondria) and is then transferred to the carboxyl residue of the c subunit. This protonation induces the transformation of the carboxyl residue to an ion-locked conformation (Pogoryelov et al., 2010) (Pogoryelov et al., 2009) and neutralizes the negative charge of the residue, allowing the c subunit to rotate apart from the a subunit toward the hydrophobic lipid layer. At the same time, the neighboring c subunit on the counterclockwise side returns from the lipid layer to form contacts with the other half channel, which has a hydrophilic environment. This contact promotes deprotonation of the carboxyl residue and induces the transformation of the c subunit to an ion-unlocked form (Symersky et al., 2012). The released proton then enters into the cytoplasmic space. Thus, proton translocation accompanies the rotation of the c-ring. For each proton, the c-ring makes one turn. Importantly, the proton translation pathway shows intrinsic chirality for the unidirectional rotation of Fo; individual protons are translated after one clockwise turn (in ATP synthesis mode) as represented as the cyan arrow in figure 8. This is highly consistent with the concept of “helical proton channel” (Minamino et al., 2008). The role of the conserved Arg in the a subunit is thought to prevent the proton shortcut without c-ring rotation (Mitome et al., 2010). In the ATP-driven proton-pumping mode, this sequence of events is reversed. The characterization of c-ring rotation against the ab2 complex in Fo is an important clue to elucidating the working principle of the Fo motor. Compared with the rotary dynamics of F1, that of Fo has been less characterized due to challenges in purifying the whole complex of FoF1 without losing any subunits (some detergents are known to promote subunit dissociation) and handling the enzyme with a highly hydrophobic part. Thus far, detergent-solubilized, fully functional FoF1 was subjected to the rotation assay under ATP hydrolysis conditions (Sambongi et al., 1999; Ishmukhametov et al., 2010; Ueno et al., 2005). The rotation of E. coli FoF1 (EFoF1) in reconstituted nanodisks was recently

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reported. In these measurements, torque-generating F1 dragged the c-ring in a counterclockwise direction. Thereby, unless the c-ring becomes a kinetic bottleneck, the dynamic features of c-ring rotation are not investigable. Only a few studies reported intervening pauses representing the multiple potential bumps that were attributed to transiently formed interaction between c-ring and the ab2 complex (Ishmukhametov et al., 2010).

REFERENCES Abrahams, J. P., Leslie, A. G., Lutter, R., Walker, J. E., 1994. Structure at 2.8 Å resolution of F1-ATPase from bovine heart mitochondria. Nature 370, 621–628. Adachi, K., Oiwa, K., Nishizaka, T., Furuike, S., Noji, H., Itoh, H., Yoshida, M., Kinosita, K., 2007. Coupling of rotation and catalysis in F-1-ATPase revealed by single-molecule imaging and manipulation. Cell 130, 309–321. Arai, H. C., Yukawa, A., Iwatate, R. J., Kamiya, M., Watanabe, R., Urano, Y., Noji, H., 2014. Torque generation mechanism of F1-ATPase upon NTP binding. Biophysical Journal 107, 156–164. Ariga, T., Muneyuki, E., Yoshida, M., 2007. F1-ATPase rotates by an asymmetric, sequential mechanism using all three catalytic subunits. Nature Structural & Molecular Biology 14, 984. Dittrich, M., Hayashi, S., Schulten, K., 2003. On the mechanism of ATP hydrolysis in F1-ATPase. Biophysical Journal 85, 2253–2266. Doering, C., Ermentrout, B., Oster, G., 1998. Rotary DNA motors. Biophysical Journal 69, 2256–2267. Guo, P., Noji, H., Yengo, C. M., Zhao, Z., Grainge, I., 2016. Biological nanomotors with revolution, linear, or rotation motion mechanism. Microbiology and Molecular Biology Reviews 80, 161–186. Hayashi, S., Ueno, H., Shaikh, A. R., Umemura, M., Kamiya, M., Ito, Y., Ikeguchi, M., Komoriya, Y., Iino, R., Noji, H., 2012. Molecular mechanism of ATP hydrolysis in F1-ATPase revealed by molecular simulations and single-molecule observations. Journal of the American Chemical Society 134, 8447–8454. Hisabori, T., Kondoh, A., Yoshida, M., 1999. The gamma subunit in chloroplast F1-ATPase can rotate in a unidirectional and counter-clockwise manner. FEBS Letters 463, 35–38. Imamura, H., Nakano, M., Noji, H., Muneyuki, E., Ohkuma, S., Yoshida, M., Yokoyama, K., 2003. Evidence for rotation of V-1-ATPase. Proceedings of the National Academy of Sciences of the United States of America 100, 2312–2315. Ishmukhametov, R., Hornung, T., Spetzler, D., Frasch, W. D., 2010. Direct observation of stepped proteolipid ring rotation in E. coli FoF1-ATP synthase. EMBO Journal 29, 3911–3923. Ito, Y., Yoshidome, T., Matubayasi, N., Kinoshita, M., Ikeguchi, M., 2013. Molecular dynamics simulations of yeast F1-ATPase before and after 16 degrees rotation of the gamma subunit. Journal of Physical Chemistry B 117, 3298–3307. Itoh, H., Takahashi, A., Adachi, K., Noji, H., Yasuda, R., Yoshida, M., Kinosita, K., 2004. Mechanically driven ATP synthesis by F1-ATPase. Nature 427, 465–468. Junge, W., Lill, H., Engelbrecht, S., 1997. ATP synthase: an electrochemical transducer with rotatory mechanics. Trends in Biochemical Sciences 22, 420–423. Kinosita, K., Jr., Yasuda, R., Noji, H., Ishiwata, S., Yoshida, M., 1998. F1-ATPase: a rotary motor made of a single molecule. Cell 93, 21–24. Kobayashi, M., Akutsu, H., Suzuki, T., Yoshida, M., Yagi, H., 2010. Analysis of the open and closed conformations of the beta subunits in thermophilic F1-ATPase by solution NMR. Journal of Molecular Biology 398, 189–199. Komoriya, Y., Ariga, T., Iino, R., Imamura, H., Okuno, D., Noji, H., 2012. Principal role of the arginine finger in rotary catalysis of F1-ATPase. Journal of Biological Chemistry 287, 15134–15142. Masaike, T., Koyama-Horibe, F., Oiwa, K., Yoshida, M., Nishizaka, T., 2008. Cooperative three-step motions in catalytic subunits of F1-ATPase correlate with 80 degrees and 40 degrees substep rotations. Nature Structural & Molecular Biology 15, 1326–1333. Meier, T., Polzer, P., Diederichs, K., Welte, W., Dimroth, P., 2005. Structure of the rotor ring of F-type Na+ATPase from Ilyobacter tartaricus. Science 308, 659–662. Menz, R. I., Walker, J. E., Leslie, A. G., 2001. Structure of bovine mitochondrial F1-ATPase with nucleotide bound to all three catalytic sites: implications for the mechanism of rotary catalysis. Cell 106, 331–341. Minagawa, Y., Ueno, H., Hara, M., Ishizuka-Katsura, Y., Ohsawa, N., Terada, T., Shirouzu, M., Yokoyama, S., Yamato, I., Muneyuki, E., Noji, H., Murata, T., Iino, R., 2013. Basic properties of rotary dynamics of the molecular motor Enterococcus hirae V-1-ATPase. Journal of Biological Chemistry 288, 32700–32707. Minamino, T., Imada, K., Namba, K., 2008. Molecular motors of the bacterial flagella. Current Opinion in Structural Biology 18, 693–701.

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Mitome, N., Ono, S., Sato, H., Suzuki, T., Sone, N., Yoshida, M., 2010. Essential arginine residue of the F0-a subunit in F0F1-ATP synthase has a role to prevent the proton shortcut without c-ring rotation in the F0 proton channel. Biochemical Journal 430, 171–177. Nadanaciva, S., Weber, J., Wilke-Mounts, S., Senior, A. E., 1999. Importance of F1-ATPase residue alphaArg-376 for catalytic transition state stabilization. Biochemistry 38, 15493–15499. Noji, H., Hasler, K., Junge, W., Kinosita, K., Jr., Yoshida, M., Engelbrecht, S., 1999. Rotation of Escherichia coli F1-ATPase. Biochemical and Biophysical Research Communication 260, 597–599. Noji, H., Yasuda, R., Yoshida, M., Kinosita, K., Jr., 1997. Direct observation of the rotation of F1-ATPase. Nature 386, 299–302. Okazaki, K., Hummer, G., 2013. Phosphate release coupled to rotary motion of F1-ATPase. Proceedings of the National Academy of Sciences of the United States of America 110, 16468–16473. Okuno, D., Iino, R., Noji, H., 2011. Rotation and structure of FoF1-ATP synthase. Journal of Biochemistry 149, 655–664. Panke, O., Cherepanov, D. A., Gumbiowski, K., Engelbrecht, S., Junge, W., 2001. Viscoelastic dynamics of actin filaments coupled to rotary F-ATPase: angular torque profile of the enzyme. Biophysical Journal 81, 1220–1233. Pogoryelov, D., Krah, A., Langer, J. D., Yildiz, O., Faraldo-Gomez, J. D., Meier, T., 2010. Microscopic rotary mechanism of ion translocation in the F0 complex of ATP synthases. Nature Chemical Biology 6, 891–899. Pogoryelov, D., Yildiz, O., Faraldo-Gomez, J. D., Meier, T., 2009. High-resolution structure of the rotor ring of a proton-dependent ATP synthase. Nature Structural & Molecular Biology 16, 1068–1088. Rees, D. M., Montgomery, M. G., Leslie, A. G. W., Walker, J. E., 2012. Structural evidence of a new catalytic intermediate in the pathway of ATP hydrolysis by F1-ATPase from bovine heart mitochondria. Proceedings of the National Academy of Sciences of the United States of America 109, 11139–11143. Rondelez, Y., Tresset, G., Nakashima, T., Kato-Yamada, Y., Fujita, H., Takeuchi, S., Noji, H., 2005. Highly coupled ATP synthesis by F1-ATPase single molecules. Nature. 433, 773–777. Sambongi, Y., Iko, Y., Tanabe, M., Omote, H., Iwamoto-Kihara, A., Ueda, I., Yanagida, T., Wada, Y., Futai, M., 1999. Mechanical rotation of the c subunit oligomer in ATP synthase (F0F1): direct observation. Science 286, 1722–1724. Shimabukuro, K., Yasuda, R., Muneyuki, E., Hara, K. Y., Kinosita, K., Yoshida, M., 2003. Catalysis and rotation of F1 motor: cleavage of ATP at the catalytic site occurs in 1 ms before 40 degrees substep rotation. Proceedings of the National Academy of Sciences of the United States of America 100, 14731–14736. Shimo-Kon, R., Muneyuki, E., Sakai, H., Adachi, K., Yoshida, M., Kinosita, K., 2010. Chemo-mechanical coupling in F1-ATPase revealed by catalytic site occupancy during catalysis. Biophysical Journal 98, 1227–1236. Spetzler, D., York, J., Daniel, D., Fromme, R., Lowry, D., Frasch, W., 2006. Microsecond time scale rotation measurements of single F1-ATPase molecules. Biochemistry 45, 3117–3124. Stracke, J. O., Fosang, A. J., Last, K., Mercuri, F. A., Pendas, A. M., Llano, E., Perris, R., Di Cesare, P. E., Murphy, G., Knauper, V., 2000. Matrix metalloproteinases 19 and 20 cleave aggrecan and cartilage oligomeric matrix protein (COMP). FEBS Letters 478, 52–56. Symersky, J., Pagadala, V., Osowski, D., Krah, A., Meier, T., Faraldo-Gomez, J. D., Mueller, D. M., 2012. Structure of the c10 ring of the yeast mitochondrial ATP synthase in the open conformation. Nature Structural & Molecular Biology 19, 485–491. Uchihashi, T., Iino, R., Ando, T., Noji, H., 2011. High-speed atomic force microscopy reveals rotary catalysis of rotorless F1-ATPase. Science 333, 755–758. Ueno, H., Suzuki, T., Kinosita, K., Yoshida, M., 2005. ATP-driven stepwise rotation of F0F1-ATP synthase. Proceedings of the National Academy of Sciences of the United States of America 102, 1333–1338. Watanabe, R., Iino, R., Noji, H., 2010. Phosphate release in F1-ATPase catalytic cycle follows ADP release. Nature Chemical Biology 6, 814–820. Watanabe, R., Matsukage, Y., Yukawa, A., Tabata, K. V., Noji, H., 2014. Robustness of the rotary catalysis mechanism of F1-ATPase. Journal of Biological Chemistry 289, 19331–19340. Watanabe, R., Okuno, D., Sakakihara, S., Shimabukuro, K., Iino, R., Yoshida, M., Noji, H., 2012. Mechanical modulation of catalytic power on F1-ATPase. Nature Chemical Biology 8, 86–92. Yasuda, R., Noji, H., Kinosita, K., Jr., Yoshida, M., 1998. F1-ATPase is a highly efficient molecular motor that rotates with discrete 120 degree steps. Cell 93, 1117–1124. Yasuda, R., Noji, H., Yoshida, M., Kinosita, K., Jr., Itoh, H., 2001. Resolution of distinct rotational substeps by submillisecond kinetic analysis of F1-ATPase. Nature 410, 898–904.

11

Mechanism of Linear Motion Peixuan Guo and Zhengyi Zhao The University of Kentucky

CONTENTS 11.1 Conserved Catalytic Cycle of Myosins................................................................................. 117 11.2 Nucleotide-Binding Region .................................................................................................. 117 11.3 Actin-Binding Region ........................................................................................................... 118 11.4 Lever Arm Region ................................................................................................................ 119 References ...................................................................................................................................... 121

11.1

CONSERVED CATALYTIC CYCLE OF MYOSINS

The modified Lymn–Taylor cycle provides the minimal framework for explaining the conserved properties of the actomyosin ATPase cycle (Guo et al., 2016; De la Cruz et al., 1999). The myosin motor domain is an ATPase that is strongly activated upon binding to actin. In the absence of any nucleotide, myosin binds to actin tightly and forms a rigor complex. ATP binding to myosin causes the cross-bridge to detach from actin and enter the weak binding states. During the detached states, ATP is hydrolyzed by myosin, and the lever arm region of myosin primes itself into a pre-power stroke state (Recovery Stroke) (Guo et al., 2016; Nesmelov et al., 2011; Trivedi, 2014). Thereafter, myosin complexed with the hydrolysis products rebinds actin in a weak binding state. This is followed by the release of Pi and ADP, which is stimulated by the binding of the complex to actin (actin-activated product release). During the actomyosin bound state, myosin pulls on the actin filament performing mechanical work, which is produced by the swing of the lever arm (Power stroke) (Guo et al., 2016; Trivedi, 2014). An additional power stroke has been shown to occur in some myosins during the ADP release step, which is thought to be associated with strain sensitivity or the ability of these mechanoenzymes to adapt to different loads. A two-ADP-state model, one with strong ADP binding affinity and the other with weak affinity, has been proposed based on kinetic, structural, and mechanical studies (Guo et al., 2016; De la Cruz & Ostap, 2004; Hannemann et al., 2005).

11.2

NUCLEOTIDE-BINDING REGION

The coordination of ATP within the nucleotide-binding pocket, cleavage of its phosphoanhydride bond, and the sequential release of products govern the mechanical cycle of myosins (Guo et al., 2016; De la Cruz & Ostap, 2004; Lymn & Taylor, 1971; Eisenberg & Greene, 1980). The ATP molecule is coordinated in the nucleotide-binding pocket (NBP)by three highly conserved structural elements, switch I, switch II, and the P-loop (Vale & Milligan, 2000; Trivedi, 2014). The P-loop NTPases family, G-proteins, kinesins, and myosins are thought to have evolved from a common ancestor. Switch I has been reported to be an important element that coordinates the sequential release of products and transmits information from the NBP to the actin-binding cleft in myosins (Guo et al., 2016; Kintses et al., 2007). Switch II is a well-conserved element that forms a salt bridge with switch I and interacts with the ɤ-phosphate of ATP which is essential for catalysis (Sasaki et al.,

DOI: 10.1201/9780429203367-12

117

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1998; Onishi et al., 2002). The P-loop is also involved in the coordination of the α and β phosphates of ATP. Additionally, magnesium (Mg) is coordinated to the oxygen on the β and ɣ phosphates and makes a direct or water-mediated contact with residues of switch I (Trivedi, 2014). The ɣ-phosphate of ATP plays a central role in the interaction of the switch elements and the P-loop with ATP. This explains why ATP and not ADP can lead to the weak actin-binding state of myosin and also induce the recovery stroke of the lever arm (Guo et al., 2016). The switch elements undergo a conformational change to a “closed” state upon binding of ATP, which leads to a twisting of a seven-stranded β sheet (transducer) and results in the opening of the actin-binding cleft (Coureux et al., 2004). Moreover, the twist of the transducer region also translates toward the C-terminal lever-arm region via a highly conserved structural element called the relay helix (Trivedi, 2014). The relay helix communication pathway induces the recovery stroke and formation of the pre-power stroke state of the lever arm. The actin-binding cleft is a deep cleft between the upper (U50) and lower (L50) 50-kDa subdomains in the motor domain (Guo et al., 2016). The binding of myosin to actin is proposed to cause movement of switch I, inducing a loss of Mg coordination and reducing its affinity and its eventual release (Rosenfeld et al., 2005). Rearrangements in the P-loop result in an isomerization to the weak ADP state of the pocket and eventual release of ADP. It has been speculated that the coupling associated with actin and ADP binding requires the coordination of bound magnesium (Hannemann et al., 2005). Moreover, two different states of switch I have been reported when MgADP is bound in the pocket, and both of these states bind ADP differently (Hannemann et al., 2005). These results suggest a role of switch I and magnesium in governing the closed–open transition of NBP and ADP release. In rapidly contracting muscle fibers, ADP release has been shown to be a major determinant of the maximum shortening velocity and has been speculated to be a central step for sensing load on the myosin cross-bridge, thus making it a strain-sensitive step (Nyitrai & Geeves, 2004). Since the lever arm senses the load, there must be allosteric communication between the lever arm and the NBP to modulate the load-dependent release of ADP (Guo et al., 2016).

11.3

ACTIN-BINDING REGION

The open–closed transition of switch I may be coupled to the closed–open equilibrium of the actinbinding cleft. A 32-amino-acid-long alpha helix that traverses the upper 50-kDa domain of myosins, called the HO helix, and a related HG/HH helix have been demonstrated by molecular modeling studies to be strongly coupled during the open–closed transition of the cleft (Guo et al., 2016; Ovchinnikov et al., 2010). It is worth noting that the structural element of switch I is located at one end of the HG/HH helix and the displacement of switch I and the HG/HH helix is proposed to cause a pull on the HO helix, which can open or close the actin-binding cleft by a tightly coupled hydrogen bonding pattern (Ovchinnikov et al., 2010). Moreover, the HO helix also makes an important connection to the switch II region, which has been shown to be important in communicating nucleotide-mediated changes to the lever arm. Conformational changes in the HO helix during the myosin ATPase cycle were examined by monitoring an endogenous tryptophan residue in smooth muscle myosin, which demonstrated that the conformational change in this helix correlates with ATP-induced dissociation and attachment to actin. However, no direct experimental studies have shown the importance of the HO helix in relation to communication with switch I and switch II. The relay helix near the lower 50-kDa domain is a 4.7-nm-long α-helix that has been well documented to be an essential feature of the force-generating region of myosin (Nesmelov et al., 2011; Agafonov et al., 2009; Muretta et al., 2013). It connects the nucleotide-binding site to the lever arm region and goes from a kinked to straight conformation during the formation of the pre-power stroke state (Nesmelov et al., 2011; Muretta et al., 2013; Kintses et al., 2008). The HO helix and relay helix are connected via the switch II loop (Guo et al., 2016). An interesting hypothesis is that the status of a nucleotide inside the pocket can be communicated to the actin-binding cleft via the HO helix and to the lever arm via the relay helix.

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11.4

119

LEVER ARM REGION

Lever arm movement during the recovery and power stroke stages of the catalytic cycle has been probed indirectly by a number of studies (Guo et al., 2016). Monitoring the intrinsic fluorescence signal originating from the tryptophan residue located at the distal end of the relay helix near the converter region has yielded insights into the coupling of the open–closed transition of switch II and the apparent position of the lever arm (Conibear et al., 2004; Malnasi-Csizmadia et al., 2000). However, studies based on tryptophan fluorescence provide no direct evidence of lever arm rotation. More recently, there have been studies measuring the lever arm swing in Dictyostelium myosin by utilizing strategically placed fluorescent resonance energy transfer (FRET)probes on the relay helix (Muretta et al., 2013). In this study, they indirectly measured the swing by correlating the kinked to the straight conformation of the relay helix to the power stroke. Another study has shown that by utilizing electron paramagnetic resonance (EPR) and transient time-resolved FRET, helix straightening occurs after actin binding and before Pi release (Muretta et al., 2013). The authors have hypothesized that relay helix straightening gates Pi release, which in turn provides the thermodynamic driving force for force generation. They have also reported that the reverse movement of the relay helix from a straight to a kinked conformation is associated with the reversal of the power stroke or the recovery stroke. The straight-to-kinked transition of the relay helix occurs after ATP binding and before hydrolysis (Guo et al., 2016). Hence, these studies report the movement of the lever arm during the recovery stroke and power stroke based on the conformation of the relay helix. Future studies that can directly measure the structural kinetics of the lever arm swing and simultaneously measure the timing of the product release steps are necessary. Modeling studies based on structural models of Dictyostelium myosin II have yielded insights into the structural mechanism of the recovery stroke (Fischer et al., 2005; Koppole et al., 2007). However, due to a lack of crystal structures in the actin-bound states, it has been difficult to perform modeling studies of the movement of the lever arm during the power stroke. Preller and Holmes (2013) have also performed targeted molecular dynamics simulations with Dictyostelium Myo II and found that soon after actin binding, a 16° rotation of the L50kDa domain puts strain on the helix that is connected to the actin-binding site. The strain twists the beta-sheet connected to this helix, which can drive the power stroke without opening switch I or switch II. They have proposed that during the power stroke, switch II moves, thus opening an exit route for Pi to escape, which would explain the actin-activated phosphate release. Several studies based on muscle fiber mechanics have given insights into the timing of the forcegenerating step in the intact sarcomere. Dantzig et al. (1992) have measured force generation and decline in tension after photolysis of caged Pi on glycerol-extracted fibers from rabbit psoas muscle. In the tension recordings, soon after Pi release, a lag of several milliseconds is observed before the force declines. The authors have proposed a two-step mechanism of force generation and Pi release with force generation preceding the release of Pi from the active site. Another report has investigated the timing of Pi binding/release and the mechanism of force generation in rabbit fasttwitch muscle fibers by employing the method of sinusoidal analysis (Kawai & Halvorson, 1991). These studies propose that a conformational isomerization precedes Pi release. Further, they also infer a distinct ADP-bound state of the cross-bridge and propose the transition between the two ADP-bound states as a rate-limiting step of the cycle. Studies performed on rabbit psoas muscles by Nagano and Yanagida (1984), Homsher and Millar (1990), and Sleep et al., (2005); on Lethocerus muscles by Molloy et al., (1987); and frog skeletal muscle fibers by Brozovich et al. (1988) also show similar results of a rapid lever arm swing before Pi release. However, laser temperature jump experiments performed on rabbit psoas muscle fibers have predicted a mechanism wherein Pi release provides the energy to generate tension, by swinging the lever arm in a force-generating state (Davis & Rodgers, 1995; Davis & Rodgers, 1995). Single-molecule studies have demonstrated the reversibility of the force-generating lever arm swing under high loads without the net utilization of ATP (Takagi et al., 2006). Sellers and Veigel

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(2010) have reported the reversibility of the power stroke with myosin V at intermediate loads (2–5 pN). However, in their system, the loads were applied at 3 ms, and considering a Pi release rate of 250 sec−1, it is highly likely that the Pi is already released when the load is applied. It is difficult to gain insights into the timing of the working stroke and the Pi release step in such a setup. A recent study by Capitanio et al. (2012) has demonstrated that a decrease in the amplitude of the working stroke in muscle fibers at high loads is due to a premature dissociation pathway that becomes more populated at higher forces. The temporal resolution of the working stroke obtained by this group is within an interval of 2 ms after the initial binding of skeletal muscle myosin to the actin filament. This provides evidence of a fast power stroke that may precede Pi release, especially in skeletal myosin where Pi release is slow, being the rate-limiting step of the catalytic cycle (45 sec−1). Debold et al. (2013) have investigated the impact of Pi on the force-generating capacity of a small ensemble of skeletal myosin molecules in an optical trap setup. They have demonstrated that in the presence of excess Pi, myosin can prematurely detach from the strongly bound state without a reversal of the power stroke. They have proposed a model wherein the power stroke is completed on actin before Pi rebinds in the AM.ADP state. This work hints at a conformational change that precedes the release of Pi. An additional swing of the lever arm is reported to occur in a number of myosins, which is associated with the actomyosin ADP state. Uemura et al. (2004) have demonstrated that the working stroke of myosin V is composed of two substeps. Two ADP-associated states are assumed in the study, which have been shown to exist with monomeric myosin V as well. They have predicted that the ADP-associated swing occurs around the time after the release of Pi and before the formation of the weak ADP-binding state. Another study has proposed a model wherein a 5-nm substep of the power stroke is accomplished by a dimeric myosin V followed by the release of ADP. This substep acts as a gate to relieve the strain that is generated by the binding of both heads to the actin filament. Biphasic working strokes are also known to occur in brush border myosin I, rat liver myosin I (Veigel et al., 1999), and smooth muscle myosin (Veigel et al., 2003). Suzuki et al. (1998) have tested the lever arm hypothesis in Dictyostelium myosin II by using FRET. They have demonstrated the reversible movement of the C-terminal fluorophore upon ATP binding and Pi release. However, this study has utilized a chimeric myosin II which has its native C-terminal lever arm replaced by a fluorescent protein. The second fluorescent protein has been fused to the N-terminus via a spacer. This study has claimed that a swing of the lever arm is observed with a rate that correlates with the release rate of Pi in Dictyostelium Myo II. However, measurements of the working stroke as a function of actin to obtain the maximal rate of the lever arm swing were not performed in this study. Moreover, a study using a similar construct that had a yellow FP (YFP)–cyan FP (CFP) fusion construct instead of the green fluorescent protein (GFP)– blue FP (BFP) construct that was used earlier has yielded different results (Zeng et al., 2006). This study has demonstrated that it is challenging to extract the precise amount of FRET in processes involving GFP-type probes because of contributions from certain non-FRET processes that can alter the donor and acceptor emission ratios. Moreover, it is challenging to measure the distance between GFP-type probes because of unknown orientation factors for these fluorophores. The use of fusion proteins can complicate FRET measurements because of their large sizes, which can affect the conformational changes of the protein under investigation. Allosteric communication pathways are at the heart of force generation in myosin motors. It is critically important to uncover these coupling pathways, which will help us understand the molecular basis of disease-causing mutations in myosin motors and may facilitate drug design paradigms to target these diseases. There are three major aspects that will greatly enhance our understanding of the allosteric communication between the nucleotide and actin-binding regions and the lever arm. First, it is important to determine whether the switch II loop is involved in coupling between the actin- and nucleotide-binding regions in Myosin V (MV). Second, it is critical to examine the role of magnesium (Mg) in mediating the conformational change of the NBP in the ADP-bound states. Lastly, a very crucial and long-standing issue in the actomyosin field is determining the precise timing of force generation and the product release steps. Investigating these aspects will shed

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light on the coupling of the mechanical and chemical steps of the catalytic cycle of myosin motors and provide novel insights into nature’s elegant design of energy transduction in molecular motors. As expected from their structural similarity, kinesin and myosin share some common features in their structural mechanism of motion generation. The conserved nucleotide-binding region of kinesin can be directly linked to the microtubule-binding and force-generating elements. The switch II region is linked to a long helix (alpha-4) that essentially functions as the relay helix connecting the microtubule and nucleotide-binding regions. The relay helix also impacts the conformation of the neck linker by altering interactions with an adjacent helix (alpha-6). The conformation of the switch I region also plays a role in microtubule binding by coordination of key microtubule-binding elements (alpha-3 and loop-8).

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Lymn, R. W., Taylor, E. W., 1971. Mechanism of adenosine triphosphate hydrolysis by actomyosin. Biochemistry 10, 4617–4624. Malnasi-Csizmadia, A., Woolley, R. J., Bagshaw, C. R., 2000. Resolution of conformational states of Dictyostelium myosin II motor domain using tryptophan (W501) mutants: implications for the openclosed transition identified by crystallography. Biochemistry 39, 16135–16146. Molloy, J. E., Kyrtatas, V., Sparrow, J. C., White, D. C. S., 1987. Kinetics of Flight Muscles from Insects with Different Wingbeat Frequencies. Nature 328, 449–451. Muretta, J. M., Petersen, K. J., Thomas, D. D., 2013. Direct real-time detection of the actin-activated power stroke within the myosin catalytic domain. Proc Natl Acad Sci U S A 110, 7211–7216. Nagano, H., Yanagida, T., 1984. Predominant attached state of myosin cross-bridges during contraction and relaxation at low ionic strength. J Mol Biol 177, 769–785. Nesmelov, Y. E., Agafonov, R. V., Negrashov, I. V., Blakely, S. E., Titus, M. A., Thomas, D. D., 2011. Structural kinetics of myosin by transient time-resolved FRET. Proc Natl Acad Sci U S A 108, 1891–1896. Nyitrai, M., Geeves, M. A., 2004. Adenosine diphosphate and strain sensitivity in myosin motors. Philos Trans R Soc Lond B Biol Sci. 359, 1867–1877. Odronitz, F., Kollmar, M., 2007. Drawing the tree of eukaryotic life based on the analysis of 2,269 manually annotated myosins from 328 species. Genome Biol 8, R196. Onishi, H., Ohki, T., Mochizuki, N., Morales, M. F., 2002. Early stages of energy transduction by myosin: roles of Arg in switch I, of Glu in switch II, and of the salt-bridge between them. Proc Natl Acad Sci U S A 99, 15339–15344. Ovchinnikov, V., Trout, B. L., Karplus, M., 2010. Mechanical coupling in myosin V: a simulation study. J Mol Biol 395, 815–833. Preller, M., Holmes, K. C., 2013. The myosin start-of-power stroke state and how actin binding drives the power stroke. Cytoskeleton 70, 651–660. Rosenfeld, S. S., Houdusse, A., Sweeney, H. L., 2005. Magnesium regulates ADP dissociation from myosin V. J Biol Chem 280, 6072–6079. Sasaki, N., Shimada, T., Sutoh, K., 1998. Mutational analysis of the switch II loop of Dictyostelium myosin II. J Biol Chem 273, 20334–20340. Sellers, J. R., Goodson, H. V., 1995. Motor proteins 2: myosin. Protein Profile 2, 1323–1423. Sellers, J. R., Veigel, C., 2010. Direct observation of the myosin-Va power stroke and its reversal. Nat Struct Mol Biol 17, 590–595. Sleep, J., Irving, M., Burton, K., 2005. The ATP hydrolysis and phosphate release steps control the time course of force development in rabbit skeletal muscle. J Physiol 563, 671–687. Suzuki, Y., Yasunaga, T., Ohkura, R., Wakabayashi, T., Sutoh, K., 1998. Swing of the lever arm of a myosin motor at the isomerization and phosphate-release steps. Nature 396, 380–383. Takagi, Y., Homsher, E. E., Goldman, Y. E., Shuman, H., 2006. Force generation in single conventional actomyosin complexes under high dynamic load. Biophys J 90, 1295–1307. Trivedi, D. V., 2014. Allosteric communication and force generation in myosin motors. Doctoral dissertation. Pennsylvania State University, University Park, PA. Uemura, S., Higuchi, H., Olivares, A. O., De la Cruz, E. M., Ishiwata, S., 2004. Mechanochemical coupling of two substeps in a single myosin V motor. Nat Struct Mol Biol 11, 877–883. Vale, R. D., Milligan, R. A., 2000. The way things move: looking under the hood of molecular motor proteins. Science. 288, 88–95. Veigel, C., Coluccio, L. M., Jontes, J. D., Sparrow, J. C., Milligan, R. A., Molloy, J. E., 1999. The motor protein myosin-I produces its working stroke in two steps. Nature 398, 530–533. 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. Nat Cell Biol 5, 980–986. Zeng, W., Seward, H. E., Malnasi-Csizmadia, A., Wakelin, S., Woolley, R. J., Cheema, G. S., Basran, J., Patel, T. R., Rowe, A. J., Bagshaw, C. R., 2006. Resonance energy transfer between green fluorescent protein variants: complexities revealed with myosin fusion proteins. Biochemistry 45, 10482–10491.

12

Finding of Widespread Viral and Bacterial Revolution dsDNA Translocation Motors Distinct from Rotation Motors by Channel Chirality and Size Gian Marco De-Donatis1, Zhengyi Zhao, and Shaoying Wang University of Kentucky

Lisa P. Huang Medical Diagnostic Laboratories, L.L.C.

Chad Schwartz, Oleg V. Tsodikov, Hui Zhang, Farzin Haque, and Peixuan Guo University of Kentucky

CONTENTS Abbreviations ................................................................................................................................. 124 12.1 Background ........................................................................................................................... 124 12.2 Results and Discussion ......................................................................................................... 125 12.2.1 Revolution and Rotation Motors Can Be Distinguished by Motor Channel Size .... 125 12.2.2 Conductance Assay of Single Connector Channels for Translocation of Tetra-Stranded DNA Reveals a Threefold Width of Phi29 Channels Compared to dsDNA ................................................................................................ 126 12.2.3 The Left-Handed Chirality of Revolution Motors Is Distinct from the Right-Handed Chirality of Rotation Motors............................................................. 127 12.2.4 Common Force Generation Mechanism of dsDNA Translocation Motors in Bacteria, Eukaryotic, and Prokaryotic Viruses ........................................................ 128 12.2.5 DNA Twists Rather Than Rotates due to Motor Channel Conformational Changes during DNA Translocation......................................................................... 130 12.2.6 Single-Molecule Real-Time Imaging and Force Spectroscopy Revealed that No Rotation Occurs during DNA Translocation ...................................................... 132 12.3 Conclusion ............................................................................................................................ 133 12.4 Materials and Methods ......................................................................................................... 134 12.4.1 Incorporation of the Connector Channel into a Planar Bilayer Lipid Membrane...... 134 12.4.2 Construction of Tetra-Stranded DNA ....................................................................... 134 12.4.3 Single-Channel Conduction Assays for Each Membrane-Inserted Connector Channel ................................................................................................... 134 12.2.4 Direct Observation of DNA Translocation ............................................................... 134 DOI: 10.1201/9780429203367-13

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Competing Interests ....................................................................................................................... 134 Authors’ Contributions .................................................................................................................. 135 Acknowledgments.......................................................................................................................... 135 Note ................................................................................................................................................135 References ...................................................................................................................................... 135

ABBREVIATIONS dsDNA, Double-stranded DNA; ASCE, Additional strand catalytic E; EMSA, Electrophoretic mobility shift assays; smFRET, Single-molecule fluorescence resonance energy transfer

12.1

BACKGROUND

Transportation of dsDNA from one cellular compartment to another is a prevalent process in all living systems. Many members of the ASCE superfamily are nanomotors with a hexameric arrangement of subunits that facilitate a wide range of functions, including dsDNA riding, tracking, packaging, and translocation, which are critical to many processes such as DNA repair, replication, recombination, chromosome segregation, transcription, and cellular reorganization [1,2]. Despite their functional diversity, a common feature of the motors of this family is their ability to convert energy obtained from the binding or hydrolysis of ATP into mechanical energy, which results in local/global protein unfolding, complex assembly/disassembly, or grabbing/ pushing dsDNA for translocation [1–11]. The hexagonal shape of the motor facilitates bottom-up assembly in nanomachine manufacturing [12–20]. Nanobiomotors have previously been classified into two main categories: linear and rotational motors, which have been clearly documented using single-molecule imaging and X-ray crystallography [21–26]. During replication, dsDNA viruses translocate their genomic DNA into pre-formed protein shells (procapsids) [27–33]. This entropically unfavorable process is accomplished by a nanomotor that uses ATP as an energy source [34–42]. This dsDNA packaging motor consists of a connector channel and packaging molecules to carry out its activities. For 35 years, it has been popularly believed that DNA packaging in dsDNA viruses involves rotation motors [43], which is seemingly supported by the swivel structure in the crystal structures of all connector channels of bacteriophages [44–46]. However, extensive investigations revealed that the dsDNA packaging motor channels do not rotate during motor actions [47–51]. For example, the T4 DNA packaging motor remains active when the motor channel protein is crosslinked to the protein shell [47]. Single-molecule imaging further verified that there is no rotation of the channel during packaging [48]. These pieces of evidence have brought up a puzzle concerning how packaging can involve a rotation motor without any rotating components. In 2010, another question was raised regarding the inverse orientations of the Phi29 motor channel and dsDNA helices [52], which further questioned the involvement of rotational motion, since the rotation mechanism of dsDNA as a bolt threading onto a motor channel as a nut requires that the threads of the bolt and the nut have the same directionality. Recently, we have discovered that the bacteriophage Phi29 dsDNA packaging motor uses a revolution mechanism without rotation, coiling, or torque forces (Figure 12.1) [9,50,53,54]. The hexameric ATPase ring exercises a force to push the dsDNA through the dodecamer channel acting as a one-way valve [9,10,52]. Observation of this revolution mechanism establishes the third class of biomotors. This finding resolves many puzzles throughout the history of long-lasting studies on the motor [9,10,55]. As the translocation of dsDNA is a ubiquitous process in living systems and motors of all dsDNA bacteriophages share some common structural and functional features, we aimed at determining whether the revolution model discovered in Phi29 can generally be applied to other DNA packaging motors. Cellular counterparts that show a strong similarity to the Phi29 viral DNA packaging motor are the FtsK and SpoIIIE family motors, featuring a hexameric motor that transports DNA and separates the intertwined lengthy genomic dsDNA during cell division or binary fission [56–63].

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FIGURE 12.1 Illustration of rotation motions and revolution motions using the Phi29 revolution motor as an example. (a) 3D structure of the Phi29 dsDNA packaging motor in a side view and top view with a pRNA hexamer derived from the crystal structure [16], and the atomic force microscopy (AFM) images of the pRNA hexamer with extended loops. (b) Illustration of rotation motors like the Earth rotates around its own axis. (c) Illustration of revolution motors like the Earth revolves around the Sun without rotation. (d) Illustration of the dsDNA revolution inside the hexameric ATPase channel. Only three of the six steps are shown. (e) Illustration of the dsDNA revolution inside the dodecameric connector channel; only four of the twelve steps are shown. Neither the channel nor the dsDNA needs to rotate during the revolution through channels (for animation, see http://nanobio. uky.edu/Connector-DNA revolution.wmv).

Unwinding of the supercoiled dsDNA resulting from rotation would lead to expensive energy consumption [64]. The revolution mechanism adopted by biological systems during evolution resembles an optimized mechanism for translocation of lengthy dsDNA genome without coiling. In this report, we analyze the motor mechanism regarding force generation of Phi29 and compare its structure and mechanism with those of DNA packaging motors of SPP1, P22, T7, HK97, mimivirus, and vaccinia virus, as well as some cellular proteins such as FtsK and SpoIIIE. We also provide a simple way to distinguish between revolution and rotation motors by channel size and chirality.

12.2 12.2.1

RESULTS AND DISCUSSION Revolution and Rotation MotoRs Can be distinguisHed by MotoR CHannel size

Previous observations that only one subunit of the hexamer binds to dsDNA at a time [8,50], as well as the cooperativity and sequential action among hexameric ATPase subunits [8], confirmed the revolution of dsDNA along the channel [50]. In this revolution process, dsDNA advances by sliding along the channel wall instead of proceeding through the center of the channel. Thus, the channel would be expected to be wider than the diameter of the dsDNA to ensure sufficient space for revolution. Inspection of the motor channel size in available crystal structures and cryogenic electron microscopy (cryo-EM) data confirmed this expectation; while the width of dsDNA is 2 nm,

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FIGURE 12.2 Comparison of the channel sizes between biomotors using rotation mechanism (a) and revolution mechanism (b). The motor channels of dsDNA phages shown in the right panel all have a channel size twice the width of dsDNA, making it impossible for a bolt and nut treading mechanism to work, thus supporting revolution rather than rotation mechanism. (PDB: RepA, 1G8Y; TrwB: 1E9R; ssoMCM, 2VL6; Rho, 3ICE; E1, 2GXA; T7-gp4D, 1E0J; FtsK, 2IUU; Phi29-gp10, 1H5W; HK97 family-portal protein, 3KDR; SPP1-gp6, 2JES; P22-gp1, 3LJ5; T4-gp17, 3EZK). The pentamer and hexamer models of T4 ATPase gp17 display a channel of 3.6 and 4.0 nm, respectively [66].

the diameters of the narrowest region of the connector channels of Phi29 [46], SPP1 [65], HK97, and the ATPase ring of T4 [66], as well as the dsDNA translocase FtsK [60] of bacteria, are all higher than 3 nm (Figure 12.2). On the other hand, the diameters of the channels of rotation motors, such as replicative DNA helicases TrwB, E1, and DnaB [67–69], are lower than 2 nm (Figure 12.2). For rotation motors, the channel would thus be expected to have a similar width as the single-stranded DNA (ssDNA) to allow for the bolt and nut threading mechanism. Nonetheless, during some processes for certain rotation motors, only one strand enters the channel, while the other remains outside [5,64,67–71]. In these situations, local unwinding fluctuations of the dsDNA might cause the separation of the two strands and facilitate the threading of the ssDNA strand into the center of the hexameric ring, as suggested by smFRET experiments [72–74]. It has been reported that the ssDNA within the channel displays an A-form helical structure [69]; thus, the channel diameter should be no larger than 2 nm to allow for contact between the DNA and the channel. The situation for branch migration is more complicated and beyond the scope of this manuscript. Overall, the above data indicate that the revolution motor can be distinguished from the rotation motor by the size of the motor channel.

12.2.2

ConduCtanCe assay oF single ConneCtoR CHannels FoR tRansloCation oF tetRa-stRanded dna Reveals a tHReeFold WidtH oF pHi29 CHannels CoMpaRed to dsdna

The channel size was further assessed by single-channel conductance assays using Phi29 connector channels as a model system. A current blockage of 32% was observed for translocation of dsDNA through the connector channel (Figure 12.3a), consistent with the ratio of the cross-sectional areas of dsDNA ((2/2)2 × 3.14 = 3.14 nm2) and channel ((3.6/2)2 × 3.14 = 10.2 nm2, 10.2 nm2 ÷ 3.14 nm2 = 32%). For tetra-stranded DNA, which was constructed by DNA nanotechnology (see Section 12.4) [75], when passing through the connector channel, a blockage of ~64% was observed (Figure 12.3b). Thus, the cross-sectional area at the narrowest region of the Phi29 connector funnel is threefold the area of the dsDNA. Such a big channel size makes it impossible for a bolt and nut tracing mechanism and makes it likely that only one ATPase subunit at a time can bind to dsDNA [8,50].

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FIGURE 12.3 Demonstration of the larger size of the Phi29 connector channel than the diameter of dsDNA as proved by single-pore translocation of double-stranded and tetra-stranded DNA by conductance assay. Current blockage of the channel by double-stranded DNA was 32% (a) and by tetra-stranded DNA was 64% (b). A purified Phi29 connector was incorporated into the lipid bilayer. Switch of voltage polarity revealed that the channel allowed only unidirectional translocation of both dsDNA (a) and tetra-stranded DNA (b).

Both dsDNA and tetra-stranded DNA show one-way translocation through the Phi29 motor channel, since the switch of the electrical polarity changed the dsDNA from passable to impassable or vice versa through the channel (Figure 12.3). One-way traffic of tetra-stranded DNA reveals that the channel does not merely serve as a pathway, it plays an active role by forming contacts with translocating double- and tetra-stranded DNA.

12.2.3

tHe leFt-Handed CHiRality oF Revolution MotoRs is distinCt RigHt-Handed CHiRality oF Rotation MotoRs

FRoM tHe

From mechanistic and physical standpoints, revolution motors depend upon a left-handed channel, while rotation motors require a right-handed channel, to match the right-handed orientation seen in both B-type DNA and A-type DNA helices. Recently, it has been reported that the anti-chiral arrangement between the Phi29 channel and the dsDNA helices facilitates the revolution of the dsDNA for unidirectional translocation during packaging [50,54]. Analysis of the crystal structures of the motor channel of SPP1 [65], T7 [76], HK97, P22 [45], and Phi29 [46] revealed that all of these motor channels display the anti-chiral arrangement between the channel and the DNA helices. The helical domains of the 12 protein subunits aligned to form the connector channels in all of these phages are tilted at 30° left-handed relative to the vertical axis of the channel, resulting in a configuration that runs anti-chiral to the right-handed dsDNA helices during packaging (Figures 12.4 and 12.5a). This structural arrangement greatly facilitates the controlled motion, supporting the conclusion that the dsDNA revolves, instead of rotating, through the connector channel without producing coiling or torsional forces while touching each of the 12 connector subunits in 12 discrete steps of 30° transitions for each contact [50]. Sequence alignments do not show apparent homology among the portal proteins of SPP1, T7, and HK97 family phages. Protein sizes also vary between different bacteriophages, ranging from 36 kDa (Phi29 gp10), 57 kDa (SPP1 gp6), 59 kDa (T7 gp8), to 94 kDa (P22 gp1) [44,76]. However, these portal proteins assemble into a propeller-like structure composed of 12 subunits with a central channel that acts as a valve for DNA translocation, and they all share very similar three-dimensional structures with several conserved regions that serve a common function in DNA packaging. Secondary structure prediction was carried out in search of structural similarities. The predicted secondary structures matched almost perfectly with the known 3D arrangements, confirming the validity of the results. Among almost all of the portal proteins, a very similar pattern of strands and helices with comparable spacing and length (Figure 12.4a) was found, particularly

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FIGURE 12.4 Secondary and crystal structures of motor channels of different bacteriophages demonstrating left-handed channel wall with a twisting of 30° leftward. (a) The 3D structures of Phi29 (P04332), HK97 (Q6NFR1), SPP1 (P54309), and P22 (P26744) were predicted using the program PredictProtein with default parameter (www.predictprotein.org), revealing that the 30° left-handed regions correlated well with their respected crystal structures (PDB: Phi29-gp10, 1H5W; HK97 family-portal protein, 3KDR; SPP1-gp6, 2JES; P22-gp1, 3LJ5). The location of the 30° left-handed tilted helix in each bacteriophage connector protein subunit is framed, which all lay at the end of the α-β motif. (b) The 30° tilt helix (red) is also shown in an external view in connector 3D structures of different bacteriophages, supporting the common mechanism that DNA revolves through the 30° tilted channel by an anti-chiral arrangement in dsDNA translocation.

a sequence of α-β-α-β-β-α stretch. Detailed analysis of quaternary structures has revealed that the 30° tilted helix exists in all portal proteins of P22, SPP1, Phi29, T7, and HK97 family phages (Figure 12.4b). Further mapping studies have revealed that the position of the 30° tilt in the quaternary structure is located at the same conserved sequence at the last alpha helix of the α-β-α-β-β-α stretch (Figure 12.4a), indicating that this 30° anti-chiral arrangement plays a critical role in dsDNA packaging as it has been conserved by evolution. As mentioned above, rotation motors should have a right-handed channel to ensure parallel threading to the right-handed DNA. Indeed, crystal structure studies of helicase–DNA complexes have verified the right-handed spiral configuration of the hexameric protein–DNA complex (Figure 12.5) [69,77,78]. In some cases, motor channels adopt right-handed chirality only when the ring is distorted while in complex with DNA, such as RecA filament [77] and DnaB, which functions in a nonplanar hexameric conformation during their movement relative to DNA [69]; otherwise, it remains as a closed symmetrical ring as observed in the absence of DNA [79]. E1 helicase also adopts a right-handed staircase pattern in the conformation of side chains when bound with DNA [80]. All of these crystallographic studies suggest that these right-handed complexes use the rotation mechanism (or a mechanism similar to a rotation mechanism for RecA, where its monomers assemble on one end of the filament and disassemble on the other). It is also possible that the gp16 ATPase in the Phi29 dsDNA packaging motor also adopts a nonplanar filament assembled from continuously spiral hexamers (or assembled from dimers) rather than a planar closed ring during the DNA packaging; however, the gp16 ring might display a left-handed configuration (Figure 12.5a).

12.2.4 CoMMon FoRCe geneRation MeCHanisM oF dsdna tRansloCation MotoRs in baCteRia, euKaRyotiC, and pRoKaRyotiC viRuses The recently discovered revolution motors use a hexameric ATPase to drive the advance of dsDNA in a sequential manner. Cellular dsDNA translocases also assemble into hexameric structures [4,5,81]. The cellular components that show the strongest similarity to phage revolution motors are found in the

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129

FIGURE 12.5 Chirality comparison of revolution and rotation motors. (a) In revolution motors, the righthanded DNA revolves within a left-handed channel, such as in the connector channels of bacteriophage Phi29 [46], P22 [45], and SPP1 [65]. (b) In rotation motors, the right-handed DNA rotates through a right-handed channel via the parallel thread, with RecA [77], DnaB [69], and E1 helicase [80] shown as examples. For E1 helicase, only the inside right-handed hairpin staircases that trace along the ssDNA are shown.

bacteria FtsK and SpoIIIE family of the ASCE DNA motor group [56–58,63]. Available pieces of evidence [58,60] lead to our hypothesis that FtsK and SpoIIIE motors also use a revolution mechanism to translocate dsDNA without rotation. Indeed, translocation of dsDNA by FtsK at a rate of 1.6–1.75 base per ATP [58,60] quantitatively agrees with the Phi29 DNA packaging motor in which each ATPase subunit uses one ATP to package 1.75 nucleotides [9,34,50,53,54]. Sequence studies of motor components of large eukaryotic dsDNA viruses, such as Acanthamoeba polyphagia mimivirus (APMV), and vaccinia viruses contain a dsDNA translocation motor that is similar to that of the FtsK-HerA superfamily [63,82,83], suggesting that these viruses also use the revolution mechanism for dsDNA packaging. Computation studies provide strong evidence that the Phi29 DNA packaging motor ATPase gp16, FtsK, and the mimivirus motor ATPase all fall into the FtsK-HerA superfamily with a configuration of a hexameric motor ring [63,82]. As shown in this report, quaternary structure analysis revealed that a left-handed, 30° tilted helix arrangement exists in the channel wall of dsDNA bacteriophages P22, SPP1, Phi29, T7, and HK97. During the revolution of the dsDNA through the channel, it advances by touching the side of the channel wall instead of proceeding through the center of the channel [50,84]. As a result, the 30° left-handed direction for each transition between two connector subunits and the 30° alteration for the dsDNA to advance 1/12 of helical pitch neutrally are observed, resulting in a zero gain, that is, no rotation occurs for the dsDNA during the translocation. The proposed model of 60° per step of the FtsK hexamer (360° ÷ 6 = 60°) [58] agrees with the finding of 30° per step within the dodecamer connector channel (360° ÷ 12 = 30°) of all dsDNA bacteriophages and 60° per steps within the Phi29 hexameric ATPase gp16 [9,50,53,54]. Channel size and chirality are key factors in the identification of translocation motor types, which can reveal the motor mechanism. The channel size is a physical confinement that can be used to distinguish revolution motors from rotation motors. As shown in

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this report, examination of the motor structures from X-ray crystallography reveals that revolution motor channels are larger than rotation motor channels. The finding that heteroduplex loop structures up to 19 bases can translocate through the phage lambda portal with the same efficiency as genome packaging [85] is another indication that the channel of lambda is wider than the dsDNA as well. Revolution motors make contact with only one strand of the dsDNA in the 5′ to 3′ direction in order to revolve along the connector channel, which has been evidenced in various motors such as Phi29 [86,87] and T4 [88]. The model that dsDNA interacts with the internal surface of the hexameric ring [50,54] is in agreement with the observation in FtsK that only one strand of the dsDNA touches the internal wall of the motor channel [57,58]. Besides, further analysis of the crystal structures of phage connectors among SPP1, P22, and Phi29 [46,54,89,90] revealed four potential-relaying electropositive lysine residues lying on the predominantly negatively charged connector channel surface. Although these four positively charged layers are nonessential for motor DNA packaging activity [90], they are reported to influence DNA translocation [90,91]. Investigations into the detailed interaction of the lysine residues with the bacteriophage genome during translocation revealed that the force generation mechanism of the relaying layers inside the channel wall altered the speed of DNA translocation, resulting in four pauses [9,54]. The interaction between these positively charged lysine rings and the negatively charged phosphate backbone of the DNA suggests that SPP1, P22, and Phi29 viral dsDNA packaging motors involve an electrostatic force in DNA translocation. Furthermore, it has been reported that the dsDNA spooling in the filled capsid is a common phenomenon in all the T7, Phi29, ε15, P22, and λ phages [92–95]. The revolution mechanism explains this spooling phenomenon. During the packaging of DNA [50,54], dsDNA will spool within the procapsid naturally as a result of the revolution process. Since rotation is not involved, no coiling is generated and no free DNA terminus is required during spooling. Initially, extra room results in a random arrangement of the entering DNA; however, toward completion of packaging, it spools tighter and tighter due to revolution, which results in a more ordered orientation of the dsDNA [92–95]. In addition, the reported revolution mechanism of phage DNA packaging motors is also consistent with recent cryo-EM imaging studies showing that the T7 dsDNA core tilts from its central axis [84].

12.2.5

dna tWists RatHeR tHan Rotates due to MotoR CHannel ConFoRMational CHanges duRing dna tRansloCation

Many connector channels of dsDNA bacteriophages (Figure 12.4) adopt a left-handed channel wall to facilitate one-way traffic during dsDNA packaging into pre-assembled protein shells [52,54]. The conformational changes of the channel have been reported to be associated with this packaging process [96,97]. Such conformational changes allow conversion of the left-handed connector after completion of DNA packaging toward the opposite configuration, thus facilitating DNA one-way ejection into host cells for infection. Indeed, three steps of conformational changes of the Phi29 connector were detected (Figure 12.6a) [97] and discovered in the DNA packaging motor of SPP1 [98]. Noticeable conformational differences between isolated Phi29 connectors and connectors in virions confirm such a structural transition after DNA packaging [96]. In the Phi29 crystal structure, the connector subunit displays a 30° left-handed tilt (Figure 12.4). However, when treated as a rigid body, the crystal structure clearly does not fit into the cryo-EM density maps, indicated by a correlation coefficient as low as 0.55. After manual adjustments, the correlation coefficient was improved to 0.70, resulting in a 10° twist of the connector toward the connector axis [96]. On the other hand, the N-terminal external region is difficult to adjust to fit as a rigid body into other parts of the connector density. It was found that the N-terminal external region underwent a significant conformational shift in the DNA-filled capsid [96]. It was concluded that angular twisting and restructuring of the connector core subunit are promoted by the interactions among Phi29 DNA and

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FIGURE 12.6 Illustration of the mechanism of dsDNA twisting during DNA packaging due to conformational changes of the Phi29 connector channel. (a) Discrete three-step conformational changes of the Phi29 connector channel were detected by single-channel conductance assay with the connector embedded in the lipid bilayer. The external view of the crystal structure of the connector channel is shown on the right. (b) The C-terminal of the connector inside the procapsid is more static than the external N-terminal. As a result, the N-terminal of the connector may shift leftward during the DNA packaging, leading to the clockwise twist of the DNA that aligns within the connector channel wall.

its structural proteins [96]. Due to the dsDNA alignment with the channel wall [9,10,50,53,54] and the relatively static C-terminal internal region, a significant conformational shift in the N-terminal external region then results in a clockwise twist of the dsDNA when viewed from the C-terminus (Figure 12.6). Recently, it has been reported that a small angular twist of 1.5° per nucleotide was observed during dsDNA packaging in Phi29 [99]. Observation of such a small angular deviation per nucleotide can be explained by these conformational changes of the connector (Figure 12.6). As evidenced above [96], if the N-terminal external region is shifted more significantly than the internal C-terminal region, a leftward twist of the DNA will occur during the revolution along the connector channel (Figure 12.6b). This is in agreement with the observed clockwise twist of 1.5° per nucleotide relative to the C-terminus of the connector [99]. The reported twist of 1.5° per nucleotide or 15.75° per helical pitch of 10.5 bp [99] during dsDNA packaging cannot be taken as a rotation mechanism in which 360° per pitch or ~34° per base pair is required. Furthermore, the reported increase in the frequency of DNA twisting per nucleotide with an increase in capsid filling is in agreement with the observation that the conformational change of the channel accelerates toward the end of the

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FIGURE 12.7 Demonstration of no DNA rotation by real-time direct observation of single-motor DNA packaging. Procapsid was immobilized in glass, and the distal end of dsDNA was tethered to a bead. DNA is packaged vertically (A) or horizontally (B) toward the slide surface (graphic is not drawn to scale). (C) The motion of the bead is tracked during DNA packaging without (a and b) and with (c and d) the addition of ATP to the sample. The motion of the bead ceased at later times only when ATP was added (c) and (d) due to the physical restriction of DNA being completely packaged. (a) and (c) show the trajectories of the bead. Different colors represent different time ranges during the translocation. (b) and (d) show the changes in beads’ travel distance versus time.

packaging process [97] (Figure 12.6a). This is logical since the dsDNA is aligned with the wall of the connector channel, and when DNA packaging is close to completion, a final confirmation will be adopted and a more obvious twisting will be observed to prepare the channel for DNA ejection toward the host infection.

12.2.6

single-MoleCule Real-tiMe iMaging and FoRCe speCtRosCopy Revealed tHat no Rotation oCCuRs duRing dna tRansloCation

In order to validate the model of revolution without the need for rotation, several single-molecule imaging experiments were carried out (Figures 12.7 and 12.8). A micrometer-sized fluorescence bead was attached to the distal end of the Phi29 genomic dsDNA. DNA translocation was directly observed in real time by single-molecule imaging microscopy to detect fluorescence images revealing the displacement of the bead [49,51]. No rotation was found in these traces (Figure 12.7). To

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FIGURE 12.8 Single-molecule polarization detection to investigate motor rotation. (a) Experiment design of single-molecule polarization detection on motor pRNA rotation during DNA packaging. The motor was stalled by γ-S-ATP, and the rotation of the pRNA ring can be excluded since no anti-correlated signals of a single Cy3 fluorophore in horizontal (H) or vertical (V) channels were observed. (b and c) Typical time trajectories of Cy3 fluorescence intensity in horizontal (black) and vertical (red) channels without (b) and with (c) the addition of ATP to restart the packaging.

exclude the possibility that the lack of rotation is a result of bond freedom between the beads and DNA or due to the difficulty in optical discrimination due to the spherical nature of the beads, a cluster of magnetic beads was attached to the end of the Phi29 DNA to generate a label with an asymmetric shape (Figure 12.7b) [49]. Experiments using different setups for DNA packaging in a vertical (Figure 12.7a) and horizontal orientation (Figure 12.7b) [49] have been repeated many times, and no rotation of DNA was observed. Polarization studies have been used to study biomotors such as T4 helicase [72]. The polarization analysis of the Phi29 DNA packaging motor did not find a rotation phenomenon either (Figure 12.8). The mechanism where no DNA rotation is required during packaging is further supported by the observation that in bacteriophage T4, both DNA ends are tethered to the portal throughout DNA packaging once the packaged DNA persistence length of about 500 bp is reached, suggesting that no rotation is needed and DNA does not get tangled up [88,100]. All these observations support a revolution mechanism for phage DNA packaging without the need for rotation.

12.3

CONCLUSION

The revolution mechanism is a common feature shared by many DNA translocation motors. Inspections of structural data from eukaryotic and prokaryotic dsDNA translocases suggest that revolution and rotation motors can be distinguished by measuring the size and chirality of the DNA translocation channel. The diameter of the channels of revolution motors is higher than 3 nm, while that of the channels of rotation motors is lower than 2 nm. Revolution motors use a left-handed channel to drive the right-handed dsDNA in an anti-chiral arrangement, while some rotation motors use parallel threads with a right-handed channel. Revolution motors hold both strands of the dsDNA within the channel, while some rotation motors hold only one strand of the DNA inside the channel [5,64,67,69–71]. Such revolution motors are void of dsDNA coiling [9,50,54,55]. A small-angle left-handed twist of dsDNA, which is aligned with the channel, takes place due to the conformational shifts of the motor channel from a left-handed configuration for DNA entry to a right-handed configuration for DNA ejection for host cell infection; however, no dsDNA rotation is required for DNA packaging.

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12.4 12.4.1

Biomotors and their Nanobiotechnology Applications

MATERIALS AND METHODS inCoRpoRation oF tHe ConneCtoR CHannel into a planaR bilayeR lipid MeMbRane

The method of inserting the connector with reconstituted liposomes into a lipid bilayer has been reported previously [101]. Briefly, a Teflon film partition (aperture 200 μm in diameter) was used to separate a bilayer lipid membrane chamber (BLM) into cis- and trans-compartments. The aperture was painted two times with 0.5 μL of 3% (w/v) DPhPC n-decane solution, and the two compartments were filled with conducting buffer (1 M NaCl or 1 M KCl, 5 mM HEPES, pH 7.4). After the formation of the lipid bilayer on the aperture, the lipid/connector complexes were added to the chamber and allowed to fuse with the planar lipid bilayer.

12.4.2

ConstRuCtion oF tetRa-stRanded dna

Five strands were custom-ordered from IDT, with the following sequences: Strand-1: 5′-CGC AGA CAT CCT GCC GTA GCC TGA GGC ACA CG-3′; Strand-2: 5′-CGT GTG CCT CAC CGA CCA ATG C-3′; Strand-3: 5′-GCA TTG GTC GGA CTG AAC AGG ACT ACG CTG GC-3′; Strand-4: 5′-GCC AGC GTA GTG GAT GTC TGC G-3′; and Strand-5: 5′-TC AGT GGC TAC GGC ACC GT-3′. The five strands were annealed in stoichiometric ratio in TMS (Tris–magnesium saline) buffer (50 mM Tris–HCl, pH8.0, 100 mM NaCl, and 10 mM MgCl2) and purified in 12% (w/v) native PAGE, following previously reported procedures [102].

12.4.3

single-CHannel ConduCtion assays FoR eaCH MeMbRane-inseRted ConneCtoR CHannel

A pair of Ag/AgCl electrodes were connected directly into the cis- and trans-compartments to measure the current traces across the lipid bilayer membrane. The current trace was recorded using an Axopatch 200B patch clamp amplifier coupled with the Axon DigiData 1322A analog–digital converter (Axon Instruments) or the BLM workstation (Warner Instruments). All voltages reported were those of the trans-compartment. Data were low-band-pass-filtered at a frequency of 1 kHz and acquired at a sampling frequency of 10–100 kHz. The Patch clamp 9.1 software (Axon Instruments) was used to collect the data, and the software Origin Pro 8.0 was used to analyze all the data.

12.2.4

diReCt obseRvation oF dna tRansloCation

The stalled packaging intermediates containing biotinylated DNA were prepared by using nonhydrolyzable γ-S-ATP [103]. The intermediates were then immobilized in perfusion chambers built from glass slides and cover slips (Figure 12.7). The 0.53-mm fluorescent streptavidin microspheres (Bangs Laboratories Inc.) were bound to the protruding, biotinylated DNA end of the intermediates. After restarting the packaging reaction by adding gp16 and ATP [103], an individual DNA packaging event was observed. Epi-illumination was used. Sequential images with 8-bit digital resolution were recorded at 1 frame per second for 600 s. The pixel resolution of the images was 0.26 mm/ pixel.

COMPETING INTERESTS Peixuan Guo is a co-founder of Kylin Therapeutics, Inc., and Biomotor and RNA Nanotech Development Co. Ltd.

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AUTHORS’ CONTRIBUTIONS GMDD, ZZ, LH, FH, and CS carried out the experiments and participated in the manuscript preparation. HZ carried out the single-molecule experiments. SW carried out the sequence alignment. OT participated in the data analysis. PG conceived the concept, designed the experiment, and wrote the manuscript. All authors read and approved the final manuscript.

ACKNOWLEDGMENTS We would like to thank Dr. Guo-Min Li for his valuable comments. The work was supported by NIH grants R01 EB012135. The content is solely the responsibility of the authors and does not necessarily represent the official views of NIH. Funding to Peixuan Guo’s Endowed Chair in Nanobiotechnology position is from the William Fairish Endowment Fund. PG is a cofounder of Kylin Therapeutics, Inc., and Biomotor and RNA Nanotech Development Co. Ltd.

NOTE 1 Gian Marco De-Donatis and Zhengyi Zhao serve as co-first author.

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13

The ATPase of the phi29 DNA Packaging Motor Is a Member of the Hexameric AAA+ Superfamily Chad Schwartz, Gian Marco De Donatis, Huaming Fang, and Peixuan Guo University of Kentucky

CONTENTS Highlights���������������������������������������������������������������������������������������������������������������������������������������141 13.1 Introduction........................................................................................................................... 142 13.2 Results.................................................................................................................................... 143 13.2.1 Phi29 DNA Packaging Motor Contains Three Coaxial Rings.................................. 143 13.2.2 Native PAGE, EMSA, and CE Reveal Hexameric ATPase....................................... 144 13.2.3 Mutations of Known Motifs Suggest that phi29 gp16 is a Member of the AAA+ Superfamily of ATPases������������������������������������������������������������������������������� 145 13.2.4 Binomial Inhibition Functional Mutant Assays Validate Hexameric ATPase.......... 146 13.3 Discussion.............................................................................................................................. 147 13.4 Materials and Methods.......................................................................................................... 149 13.4.1 Cloning, Mutagenesis and Protein Purification......................................................... 149 13.4.2 Measurement of gp16 ATPase Activity..................................................................... 149 13.4.3 In Vitro Virion Assembly Assay................................................................................ 149 13.4.4 Statistical Analysis and Data Plotting....................................................................... 150 13.4.5 CE Experiments to Determine Ratio of gp16 to Bound dsDNA............................... 150 13.4.6 Native PAGE of eGFP-gp16....................................................................................... 150 13.4.7 Atomic Force Microscopy (AFM) Imaging.............................................................. 150 13.4.8 Electrophoretic Mobility Shift Assay (EMSA)......................................................... 150 Acknowledgements......................................................................................................................... 151 References....................................................................................................................................... 151

HIGHLIGHTS • • • •

Phi29 motor ATPase gp16 is a hexamer. Phi29 motor ATPase is a member of the hexameric AAA+ superfamily. Native gel reveals six gp16 bands, and increasing concentration drives gp16 to the hexamer. Binomial distribution assay confirms the presence of six copies of the ATPase on the motor. • Capillary electrophoresis confirmed that the ratio of dsDNA to ATPase gp16 is 1:6.

DOI: 10.1201/9780429203367-14

141

142

13.1

Biomotors and their Nanobiotechnology Applications

INTRODUCTION

The superfamily of AAA+ motors (ATPases Associated with diverse cellular Activities) plays a key role in several assorted functions, and many members of this clade of ATPases often fold into hexameric structures (Mueller-Cajar, Stotz et al., 2011; Wang, Mei et al., 2011). Despite their diversity, the common characteristic of this family is their ability to convert chemical energy from the hydrolysis of the γ-phosphate bond of ATP into a protein conformational change. This conformational change generates a loss of affinity for its substrate and exerts a mechanical movement, which in turn is used to either make or break contacts between macromolecules, resulting in local or global protein unfolding, assembly or disassembly of complexes, or transport of macromolecules relative to each other. These activities underlie processes critical to DNA repair, replication, recombination, chromosome segregation, dsDNA transportation, membrane sorting, cellular reorganization, and many others (Martin, Baker et al., 2005; Ammelburg, Frickey et al., 2006). dsDNA viruses package their DNA genome into a preformed protein shell called a procapsid, with the aid of a nanomotor (Feiss & Rao, 2012; Guo & Lee, 2007; Fang, Jing et al., 2012; Zhang, Schwartz et al., 2012). Since 1978, it has been popularly believed that viral DNA packaging motors run through a fivefold/sixfold mismatch rotation mechanism (Hendrix, 1978). An RNA component (pRNA) was discovered on the phi29 DNA packaging motor (Guo, Erickson et al., 1987), and subsequently, pRNA was determined to exist as a hexameric ring (Guo, Zhang et al., 1998; Zhang, Lemieux et al., 1998). Based on this structure, it was proposed that the mechanism of the phi29 viral DNA packaging motor is similar to that used by other hexameric DNA tracking motors of the AAA+ family of proteins (Guo, Zhang et al., 1998). A debate subsequently developed concerning whether the RNA and ATPase of the motor exist as hexamers (Zhang, Endrizzi et al., 2012; Guo, Zhang et al., 1998; Zhang, Schwartz et al., 2012; Shu, Zhang et al., 2007; Xiao, Zhang et al., 2008; Zhang, Lemieux et al., 1998; Ibarra, Caston et al., 2000) or as pentamers (Moffitt, Chemla et al., 2009; Yu, Moffitt et al., 2010). The differing viewpoints have not yet been fully reconciled, but we have recently shown using X-ray diffraction, atomic force microscopy (AFM) imaging, and single-molecule studies that the motor consists of three coaxial rings geared by hexameric pRNA (Zhang, Endrizzi et al., 2012) (Figure 13.1). The force generation mechanism of the phi29 DNA packaging motor is still debated (Moffitt, Chemla et al., 2009; Aathavan, Politzer et al., 2009; Jing, Haque et al., 2010; Zhang, Schwartz et al., 2012; Geng, Fang et al., 2011; Fang, Jing et al., 2012; Schwartz, Fang et al., 2012). The phi29 DNA packaging motor reconstituted in the defined system more than 20 years ago (Guo, Grimes et al., 1986) is one of the most well-studied biomotor systems and has also proven to be one of the most powerful molecular motors (Smith, Tans et al., 2001; Rickgauer, Fuller et al., 2008), capable of generating forces up to 57–110 pN. The DNA packaging mechanism has been studied extensively (Simpson, Tao et al., 2000; Aathavan, Politzer et al., 2009; Smith, Tans et al., 2001; Earnshaw & Casjens, 1980; Hendrix, 1998; Johnson, Bai et al., 2007; Rao & Black, 1985; Guo, Zhang et al., 1998; Sun, Kondabagil et al., 2007; Zhang, Lemieux et al., 1998; Zheng, Olia et al., 2008; Agirrezabala, Martin-Benito et al., 2005; Butcher, Bamford et al., 1995; Dubé, Tavares et al., 1993; Gutierrez, Freire et al., 1994; Lebedev, Krause et al., 2007; Orlova, Gowen et al., 2003; Shu, Zhang et al., 2007; Sousa & Padilla, 1995; Stewart, Fuller et al., 1993; Xiang, Morais et al., 2006; Hugel, Michaelis et al., 2007). The motor is composed of a dodecameric connector at the vertex of the procapsid, geared by a pRNA ring (Guo, Erickson et al., 1987), which encircles the N-terminus of the connector (Xiao, Moll et al., 2005; Atz, Ma et al., 2007; Xiao, Zhang et al., 2008), and a ring of gp16, which functions as an ATPase to drive the motor (Guo, Peterson et al., 1987b; Ibarra, Valpuesta et al., 2001). The connector was recently revealed to allow only unidirectional movement of dsDNA (Jing, Haque et al., 2010), and a model using a “push through a one-way valve” mechanism was described (Schwartz, Fang et al., 2012; Fang, Jing et al., 2012), which agrees well with the previously proposed ratchet (Serwer, 2003) and compression (Ray, Sabanayagam et al., 2010; Ray, Ma et al., 2010) models. This mechanism describes dsDNA as being pushed through the connector channel by the ATPase gp16, while the connector functions like a valve to prevent

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FIGURE 13.1 Depiction of the phi29 DNA packaging motor structure and function. A schematic of hexameric pRNA (left, top) and AFM images of loop-extended hexameric pRNA (top, right) {7936}. Illustrations of the phi29 DNA packaging motor and a pRNA hexamer: side view (bottom, left) and bottom view (bottom, right).

DNA from slipping out of the capsid during the packaging process (Black, 1989; Feiss & Rao, 2012; Casjens, 2011; Guo & Lee, 2007). This entropically unfavorable process is accomplished using ATP as an energy source. The ATPase gp16 is the most critical part of the phi29 DNA packaging motor. It provides energy for the motor by hydrolyzing ATP, converting energy obtained from breaking a chemical bond into physical motion. This enzyme possesses the typical Walker A and Walker B motifs (Guo, Peterson et al., 1987b) as found in many other well-characterized AAA+ proteins (Burroughs, Iyer et al., 2007; Iyer, Makarova et al., 2004). The protein has been shown to bind to the 5′/3′ paired helical region of pRNA (Lee & Guo, 2006; Koti, Morais et al., 2008), and furthermore, its ATPase activity could be stimulated by both pRNA and DNA (Guo, Peterson et al., 1987b; Lee, Zhang et al., 2008; Ibarra, Valpuesta et al., 2001; Grimes & Anderson, 1990). Intermediates in DNA packaging have been isolated (Guo, Peterson et al., 1987a; Smith, Tans et al., 2001; Koti, Morais et al., 2008; Shu & Guo, 2003), and models of gp16 supercoiling dsDNA have been proposed (Grimes & Anderson, 1997; Koti, Morais et al., 2008). Here, the oligomeric state of the ATPase has been extensively investigated in order to better understand the DNA translocation mechanism. We conclusively determined that the motor ATPase forms a hexamer in a concentration-dependent manner and upon binding to its substrate dsDNA. Furthermore, the major motifs of the ATPase have now been identified, and we have shown through mutation analysis that the phi29 ATPase is a member of the hexameric AAA+ superfamily.

13.2 13.2.1

RESULTS pHi29 dna paCKaging MotoR Contains tHRee Coaxial Rings

The phi29 DNA packaging motor consists of three major structural components: the connector, pRNA, and ATPase gp16 (Figure 13.1). Extensive studies (Guo, 2002; Green, Wang et al., 2010; Ibarra, Caston et al., 2000; Xiao, Zhang et al., 2008; Zhang, Lemieux et al., 1998; Shu, Zhang et al.,

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2007; Trottier & Guo, 1997; Chen, Trottier et al., 1997) of the pRNA and a recent crystal structure (Zhang, Endrizzi et al., 2012) have revealed that pRNA exists as a hexamer, as also confirmed by AFM (Shu, Haque et al., 2013). These data show that the three coaxial rings are connected to each other with fixed stoichiometry.

13.2.2

native page, eMsa, and Ce Reveal HexaMeRiC atpase

Fusion of eGFP to the N-terminus of gp16 results in fluorescent gp16 (eGFP-gp16) that shows biological activity similar to that of native gp16 (Lee, Zhang et al., 2009). eGFP-gp16 yields six distinct fluorescent bands on a native PAGE gel, which separates solely on the basis of mass (see Section 13.4), indicative of six monomers oligomerizing to form a hexamer (Figure 13.2a). The monomer and all even-numbered oligomer bands have a higher intensity than the trimer and pentamer, suggesting that the assembly sequence is monomer to dimer, to tetramer, and finally to hexamer, such that the final gp16 oligomeric state is likely a trimer of dimers, as in other ATPases (Sim, Ozgur et al., 2008; Skordalakes & Berger, 2006; Ziegelin, Niedenzu et al., 2003; Sim, Ozgur et al., 2008). In addition, as the concentration of gp16 increased, the intensity of the hexamer band increased significantly, while the intensity of smaller oligomers remained fairly constant, further suggesting that a hexamer is the final oligomeric state. Finally, the presence of eGFP-gp16 hexamer was further confirmed by stoichiometric ratio assays as discussed in the following sections. Electrophoretic mobility shift assays (EMSA) were employed with the fluorescent eGFP-gp16 and with a short 40-bp dsDNA fragment conjugated to a Cy3 fluorophore. The two components were mixed together with ATP and a nonhydrolyzable ATP analog (γ-S-ATP) (Figure 13.2b). The ATPase bound more tightly to the dsDNA upon the addition of γ-S-ATP (Figure 13.2b, lane 6) as observed previously (Schwartz, Fang et al., 2012). Furthermore, after the addition of ATP to the gp16–DNA complex, two distinct ATPase bands were present (Figure 13.2b, lanes 7,8), perhaps representative of two different conformational states of gp16. We repeated the EMSA with increasing amounts of ATPase and a fixed amount of dsDNA to determine the stoichiometry of the ATPase bound to dsDNA. As the molar concentration ratio of gp16 to dsDNA reached 6:1, free dsDNA (bottom band, Figure 13.3a Cy3 channel) shifted nearly entirely to the bound state (top yellow band, lane 6). Capillary electrophoresis was used to validate the qualitative EMSA data. In this case, the amount of gp16 was held at 3 µM, mainly due to the stickiness of the protein in the small capillary, and the [dsDNA] was varied in the reaction mixture.

FIGURE 13.2 (a) 6% native PAGE using a nondenaturing detergent that fractionates by size reveals distinct bands characteristic of six oligomeric states; the top, hexameric band increases as the concentration of protein is increased (15, 17.5, 20 µM). Oligomeric states were assigned based on the mobility of marker proteins in the Native PAGE Mark kit. (b) EMSA of native eGFP-gp16 (3 µM) with short 40-bp Cy3–dsDNA (300 nM) and ATP (30 mM) or γ-S-ATP (1.25 mM). The GFP channel (left) shows the migration of the ATPase, whereas the Cy3 channel (right) indicates the migration of the dsDNA. Two distinct states of ATPase exist after the addition of ATP to the gp16–DNA complex.

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FIGURE 13.3  ATPase gp16 binds to DNA in a 6:1 molar ratio. EMSA of 3 µM gp16 and dsDNA (a) where the free dsDNA band disappears (bottom right) as the molar ratio of gp16 to dsDNA reaches 6:1. (b) Capillary electrophoresis of eGFP-gp16 and Cy3–DNA complexes after quantification of fluorescent peaks. Data are plotted as a ratio of total DNA versus bound DNA and plateaus at 0.5 µM, a concentration six times less than the fixed molar concentration of ATPase gp16.

The fluorescent peak corresponding to the DNA–protein complex was quantified over a range of dsDNA concentrations. A plateau was achieved at 0.5 µM DNA bound, representing a DNA–protein ratio of 1:6, further arguing that the gp16 ATPase is a hexamer (Figure 13.3b).

13.2.3 Mutations of Known Motifs Suggest that phi29 gp16 is a Member of the AAA+ Superfamily of ATPases gp16 contains well-conserved motifs responsible for ATP binding (Walker A and arginine finger) and ATP hydrolysis (Walker B), typical of all AAA+ proteins. The Walker A motif was previously identified, but the Walker B motif was not determined {131}. Sequence alignment with AAA+ proteins revealed the Walker B motif (hhhhDE) at residues 114–119 (TIVFDE). To confirm the results of sequence alignment, relevant amino acids of both Walker A and Walker B motifs were mutated. For the Walker A motif, the previous mutant G27D was cloned. In the Walker B motif, two mutants were generated: E119A and D118E/E119D double mutant. The most important residues in Walker B were the aspartate (D) for its role in magnesium ion binding and glutamate (E) responsible for the activation of a water molecule to perform a nucleophilic attack on the gamma phosphate of ATP. Both mutants were tested for their ability to hydrolyze ATP and to bind DNA. Both mutants were subjected to the ATP hydrolysis assay (Lee, Zhang et al., 2008). Only the wildtype ATPase hydrolyzed ATP (Figure 13.4a); the Walker A G27D mutant was incapable of binding ATP, while the Walker B mutant can bind, but cannot hydrolyze. We expanded our testing of the mutants in terms of DNA binding. Using the same capillary electrophoresis assay used for wildtype ATPase, we quantified the DNA-bound peaks of both mutants. In the presence of γ-S-ATP, the wildtype and Walker B mutant displayed similar DNA binding affinities. However, upon the addition of ATP, the wildtype no longer remained bound to DNA as previously shown (Schwartz, Fang et al., 2012), but the Walker B D118E/E119D mutant retained its DNA binding capability, suggesting that this identified motif is in fact responsible for the catalytic step that pushes dsDNA away from gp16 upon hydrolysis. Lastly, we attempted to validate our findings using EMSA (Figure 13.4c). Gp16 ATPase and fluorescent DNA were mixed together and incubated at room temperature for 20 minutes. The samples were then loaded into an agarose gel. The top green gel represents the fluorescent signal of the eGFP-conjugated ATPase; however, the bottom, yellow gel shows the migration of the Cy3–fluorescent dsDNA. In the Cy3 gel, the upper bands are representative of DNA bound to gp16 ATPase as the protein retards the migration of the short DNA. However, the bottom bands are free DNA

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FIGURE 13.4 ATPase gp16 contains a Walker A and a Walker B motif typical of the AAA+ family. Assay of gp16 ATPase activity was described previously (Lee, Zhang et al., 2008) (a). Walker A G27D and Walker B D118E/E119D mutants of gp16 prevent ATP hydrolysis. Capillary electrophoresis quantification of dsDNA binding to mutant and wildtype gp16 (b). Walker B D118E/E119D mutant retains the binding capability to dsDNA despite the addition of ATP. EMSA of mutant and wildtype ATPase (c). DNA binding is diminished with the Walker A G27D mutant, but is retained in the Walker B D118E/E119D mutant with the addition of ATP or γ-S-ATP. The results were comparable with the Walker B E119A mutant.

as the negatively charged strand of nucleotides quickly migrates to the positive electrode. Again, the wildtype gp16 ATPase exhibits high affinity to dsDNA with the addition of γ-S-ATP (lane 3), but diminished affinity with ATP or no phosphate analog (lanes 2,4). The Walker A G27D mutant has diminished binding affinity in all cases (lanes 5–7), albeit higher affinity with the addition of γ-S-ATP, as this mutant is incapable of binding ATP, which stabilizes the interaction between gp16 and dsDNA. Finally, the Walker B D118E/E119D mutant, which previously has been shown to be incapable of hydrolyzing ATP, was incapable of binding without ATP (lane 8), but exhibited high affinity with both ATP and γ-S-ATP (lanes 9,10). Both the capillary electrophoresis quantification and the EMSA confirmed our hypothesis that the recently discovered Walker B motif of phi29 ATPase is responsible for ATP hydrolysis.

13.2.4

binoMial inHibition FunCtional Mutant assays validate HexaMeRiC atpase

We further demonstrated that hexameric gp16 was active in phi29 DNA packaging using a Walker B mutant gp16 and a binomial distribution analysis (Trottier & Guo, 1997; Chen, Trottier et al., 1997). The Walker B D118E/E119D mutant gp16 is completely inactive in DNA packaging. The mutant protein was mixed with wildtype in different ratios ranging from 10% to 90% at limiting quantities,

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The ATPase of the phi29 DNA Packaging Motor

FIGURE 13.5 Viral assembly inhibition assay using a binomial distribution revealing that gp16 possesses a sixfold symmetry in the DNA packaging motor (Trottier & Guo, 1997). Theoretical plot of percent Walker B mutant gp16 versus yield of infectious virions in in vitro phage assembly assays. Predictions were made with the followZ  Z  Z  Z Z   Z Z  Z −1 Z Z−M M ing equation: ( p + q) Z =   p Z +   p Z −1q +   p Z − 2q 2 +  +  q a,  pq +   q =  p M  Z 0  1  2   Z − 1 M −0  where p is the percentage of wildtype eGFP-gp16; q is the percentage of eGFP-gp16/ED; Z is the total number of eGFP-gp16 per procapsid or gp3-DNA; M is the number of mutant eGFP-gp16 in the phi29 DNA packaging motor; and p + q = 1 (Trottier & Guo, 1997).

∑

and the activity of the complex was assayed using the in vitro viral assembly system (Figure 13.5) (Lee & Guo, 1994). In this experiment, we assume that both the mutant and wildtype have an equal chance of incorporation within the final oligomer for packaging. The dominant inhibitory activity of the Walker B mutant allowed an independent means of determining the stoichiometry of the ATPase (Trottier & Guo, 1997). In these trials, we assumed that the stoichiometry, Z, of the ATPase gp16 in the complex lies between 1 and 12. Different concentrations of wildtype gp16 were mixed with the inactive Walker B mutant and used for in vitro assembly reactions. We used a binomial distribution of (p + q)Z , where p and q represent the ratio of wildtype and mutant subunits within the gp16 oligomer, respectively (Trottier & Guo, 1997). Following the expansion of the binomial, we generated 12 theoretical curves corresponding to a stoichiometry of 1–12 using a plot of motor activity (in this case, production of phi29 virions) against the ratio of the Walker B mutant. The empirical data almost perfectly overlap with the theoretical curve in slope and shape representative of a stoichiometry of 6, thereby confirming that the motor complex is hexameric (Figure 13.6).

13.3

DISCUSSION

For many years, there has been a debate as to the stoichiometry of phi29 motor components. Convincing data have been shown by both camps as to the fivefold and sixfold nature of the gp16 ATPase and packaging RNA. However, both sides are in agreement that the stoichiometries of these two components exist in a 1:1 ratio. It has previously been shown that the symmetry remains uniform between the ATPase and pRNA irrespective of whether it exists as a pentameric pRNA and pentameric ATPase or hexameric pRNA and hexameric ATPase, suggesting that the two work in unison independent of the stoichiometry. The data shown here indicate that gp16 ATPase is a member of the AAA+ superfamily of proteins, and similar to this family, the phi29 motor ATPase also exists in either a high or low-affinity state for the DNA substrate. Recently, it has been qualitatively demonstrated via EMSA (Schwartz,

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FIGURE 13.6 Hexameric push through a one-way valve mechanism (Schwartz, Fang et al., 2012). A conformational change in the hexameric ATPase occurs subsequently after binding to ATP, which confers an increase in binding affinity to dsDNA. The release of inorganic phosphate from the ATPase complex results in a power stroke to push the genomic dsDNA through the one-way valve of the connector portal protein into the capsid shell.

Fang et al., 2012) that the ATPase gp16 is capable of binding to dsDNA in the presence of γ-SATP. Fusion of a fluorescent tag on the ATPase did not affect its function or activity (Lee, Zhang et al., 2009), but provided a marker for binding assays. In the previous reports, a small amount of Cy3–dsDNA was bound by eGFP-gp16 using the EMSA. However, stronger binding of gp16 to dsDNA was observed when gp16 was incubated with γ-S-ATP and dsDNA (Schwartz, Fang et al., 2012). Also in the previous reports, Forster resonance energy transfer (FRET) analysis and sucrose sedimentation studies further validated our finding that the gp16–dsDNA complex is stabilized by the addition of γ-S-ATP {7100}. Furthermore, the data confirmed that gp16 possesses both a DNA binding domain and a Walker A motif with which to bind ATP (Schwartz, Fang et al., 2012). By sequence homology and point mutation analysis, this motif has now been shown to be responsible for ATP hydrolysis in the ATPase of phi29. As expected, all the mutants were severely impaired in ATP hydrolysis activity and were similar to the Walker A mutant G27D, proving that the Walker A motif is responsible for the binding of ATP. Regarding the ability to bind to DNA in the presence of γ-S-ATP, mutations in the walker A motif displayed a limited ability to bind DNA compared with the wildtype (Figure 13.4b and c), most likely due to their impaired affinity for γ-S-ATP. On the contrary, the walker B mutants retained their binding affinity for DNA in the presence of γ-S-ATP and were also sufficient to bind DNA in the presence of ATP, confirming that the Walker B mutation only affects the ability to hydrolyze ATP, but not the binding to the nucleotide. Our data show that in the absence of ATP, or its derivative γ-S-ATP, the binding of gp16 to DNA is reduced. However, after the addition of γ-S-ATP, the binding efficiency of gp16 to DNA increased significantly (Figure 13.4b and c). This suggests that ATP induces a change in gp16 that causes it to assume a high-affinity conformation for dsDNA binding, a conclusion strengthened by the inability of the Walker A mutant protein, which cannot bind ATP, to elicit a conformational change. In the previous report, when ATP was added to the gp16–γ-S-ATP–dsDNA complex, rapid ATP hydrolysis was observed (Schwartz, Fang et al., 2012) and gp16 dissociated from the dsDNA. This indicates that after hydrolysis, gp16 undergoes a further conformational change that produces an external force against the dsDNA that pushes the substrate away from the motor complex by a power stroke. This phenomenon can be seen in Figure 13.2b in which the ATPase exists in two states after the addition of ATP: DNA bound or expelled. However, introducing a mutation to the Walker B motif eliminated the catalytic force step. These data correlate nicely with other reports that Walker B mutants do not hydrolyze ATP, but bind strongly to DNA.

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Gp16 is a DNA-dependent ATPase of the phi29 DNA packaging motor (Guo, Peterson et al., 1987b; Ibarra, Valpuesta et al., 2001; Lee, Zhang et al., 2008). Energy is provided to the motor through ATP. As mentioned previously, non-hydrolyzable γ-S-ATP stalled and fastened the gp16– dsDNA complex. It has been found that the hydrolysis of ATP leads to the release of dsDNA from gp16. After ATP was added to the gp16–dsDNA–γ-S-ATP complex, the band representing the gp16–dsDNA complex disappeared (Schwartz, Fang et al., 2012). The release of dsDNA from the gp16–dsDNA–γ-S-ATP complex by ATP was also demonstrated by sucrose gradient sedimentation {7100}. Hydrolysis of ATP was confirmed when the purified gp16–dsDNA–γ-S-ATP hydrolyzed ATP after the addition of ATP to the purified complex {7100}. These results suggested that hydrolysis of ATP leads to the release of dsDNA from the gp16, forcing the DNA substrate away from the interior pocket of the ATPase and lending to the physical motion of the genomic DNA toward the capsid. Our data combining the stoichiometry of the ATPase and the sequential action previously elucidated (Schwartz, Fang et al., 2012) allow us to build upon our previous “push through a oneway valve” DNA packaging model. After binding to ATP, the ATPase undergoes a conformational change which significantly increases its affinity to dsDNA. An additional conformational change of the ATPase after the release of inorganic phosphate causes gp16 to perform a power stroke to push dsDNA into the portal protein. The stoichiometry of the phi29 DNA packaging motor has long been a contentious subject. Here, we have provided additional biochemical data showing that the ATPase gp16 consists of six subunits (Figure 13.2a), upon binding to dsDNA (Figure 13.3), and also in the active phi29 motor (Figure 13.5). Furthermore, we have identified the classical Walker motifs typical of the hexameric AAA+ superfamily and found that the phi29 DNA packaging motor uses a revolution without rotation and coiling or generation of torque (Schwartz, Zhang et al., 2013) (accompanying paper). In our accompanying paper, we show that the ATPase “hands off” the substrate dsDNA in a sequential action manner lending to revolution around the ATPase and connector protein. Our data lead to the conclusion that the hexameric stoichiometry and the mechanism of revolution for phi29 DNA packaging motor are in accordance with FtsK of the hexameric AAA+ superfamily, and we expect that most phages follow this “push through a one-way valve” via revolution mechanism (Zhao, Khisamutdinov et al., 2013; Schwartz, Zhang et al., 2013) (accompanying paper).

13.4 MATERIALS AND METHODS 13.4.1

Cloning, Mutagenesis and pRotein puRiFiCation

The engineering of eGFP-gp16 and the purification of the gp16 fusion protein have been reported previously (Lee, Zhang et al., 2009). The eGFP-gp16 mutants G27D, E119A, and D118E E119D were constructed by introducing mutations in the gp16 gene (Keyclone Technologies).

13.4.2

MeasuReMent oF gp16 atpase aCtivity

Enzymatic activity via fluorescent labelling was described previously (Lee, Zhang et al., 2008). Briefly, a phosphate-binding protein conjugated to a fluorescent probe that senses the binding of phosphate was used to assay ATP hydrolysis.

13.4.3

In VItro viRion asseMbly assay

Purified in vitro components were mixed and were subjected to the virion assembly assay as previously described (Lee & Guo, 1994). Briefly, newly assembled infectious virions were inoculated with Bacillus bacteria and plated. The activity was expressed as the number of plaques formed per volume of sample (pfu/mL).

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13.4.4

Biomotors and their Nanobiotechnology Applications

statistiCal analysis and data plotting

Most of the statistical analyses were performed using Sigmaplot 11. The Hill coefficient was determined by nonlinear regression fitting of the experimental data to the following equation: E = Emax*(x)n / (kapp +(x)n), where E and Emax refer to the concentration of the gp16–DNA complex, x is the concentration of ATP or ADP, kapp is the apparent binding constant, and n is the Hill coefficient.

13.4.5

Ce expeRiMents to deteRMine Ratio oF gp16 to bound dsdna

CE (capillary electrophoresis) experiments were performed on a Beckman MDQ system equipped with double fluorescence detectors (at 488 nm and 635 nm excitation wavelength). A bare borosilicate capillary with a total length of 60 cm and a 50 µm inner diameter was used. Assay conditions contained separation buffer of 50 mM Tris–HCl, 100 mM sodium borate at pH 8.00, 5 mM MgCl2, 10% PEG 8000 (w/v), 0.5% acetone (v/v), 3 mM eGFP-gp16 monomer, and variable amounts of ATP/ADP and DNA.

13.4.6

native page oF egFp-gp16

Increasing amounts of eGFP-gp16 were loaded onto a 6% Tris–glycine polyacrylamide gel in conjunction with the Native PAGE Mark kit (Invitrogen). This commercially available Native PAGE Mark uses a nondenaturing detergent to mildly solubilize and coats the protein with a negative charge. Thus, gel electrophoresis separates solely on the basis of mass. The gel was imaged using a Typhoon gel image scanner at an excitation wavelength of 488 nm.

13.4.7

atoMiC FoRCe MiCRosCopy (aFM) iMaging

APS mica was obtained by incubation of freshly cleaved mica in 167 nM 1-(3-aminopropyl) silatrane as described (Shlyakhtenko, Gall et al., 2003; Lyubchenko & Shlyakhtenko, 2009). Native PAGE-purified RNA samples were diluted with 1xTMS buffer to a final concentration of 3–5 nM. Then, 5–10 µL of pRNA was immediately deposited on the APS mica surface. After 2 minutes of incubation, excess samples were washed with diethyl pyrocarbonate (DEPC)-treated water and dried under a flow of Argon gas. AFM images in air were acquired using the MultiMode AFM NanoScope IV system (Veeco/Digital Instruments, Santa Barbara, CA) operating in the tapping mode. Two types of AFM probes were used under tapping mode imaging in air: (i) regular tapping mode silicon probes (Olympus from Asylum Research, Santa_Barbara, CA) with a spring constant of ~42 N/m and a resonant frequency between 300 and 320 kHz and (ii) noncontact NSG01_DLC probes (K-Tek Nanotechnology, Wilsonville, OR) with a spring constant of about 5.5 N/m and a resonance frequency between 120 and 150 kHz.

13.4.8

eleCtRopHoRetiC Mobility sHiFt assay (eMsa)

The fluorescently tagged protein that facilitates detection and purification was shown to possess similar assembly and packaging activity as compared to the wildtype (Lee, Zhang et al., 2009; Schwartz, Fang et al., 2012). Cy3–dsDNA (40 bp) was prepared by annealing two complementary DNA oligos containing a Cy3 label (IDT) at its 5′ ends and purified by a 10% polyacrylamide gel. Samples were prepared in 20 μL buffer A (20 mM Tris–HCl, 50 mM NaCl, 1.5% glycerol, 0.1 mM Mg2+). Specifically, 1.78 μM eGFP-gp16 was mixed with 7.5ng/μL of 40 bp Cy3–DNA in the presence or absence of ATP and γ-S-ATP. The samples were incubated at ambient temperature for 20 minutes and then loaded onto a 1% agarose gel (44.5 mM Tris and 44.5 mM boric acid) and electrophoresed at 4°C for 1 hour at 8 V/cm. The eGFP-gp16 and Cy3–DNA samples were analyzed by a fluorescent LightTools Whole Body Imager using 488 nm and 540 nm excitation wavelengths for GFP and Cy3, respectively.

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ACKNOWLEDGEMENTS We would like to thank Dr. Guo-Min Li for his valuable comments; Yi Shu, Luda Shlyakhtenko, and Yuri Lyubchenko for the AFM images of pRNA hexamer; Zhengyi Zhao, Emil Khisamutdinov, and Hui Li for their diligent work on the animation figures; and Jeannie Haak for editing this manuscript. The work was supported by NIH grants R01 EB012135 and U01 CA151648 to PG, who is a co-founder of Kylin Therapeutics, Inc, and Biomotor and Nucleic Acids Nanotech Development, Ltd.

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Arginine Finger Serving as the Starter of Viral DNA Packaging Motors Chenxi Liang, Chun Chan, Zhefeng Li, Xiaolin Cheng, and Peixuan Guo The Ohio State University

CONTENT References ...................................................................................................................................... 156 Generally speaking, in a car engine, the starter is the part to initiate the motor operation. Biological motors are nanoscale machines ubiquitous in many biological processes in both prokaryotes and eukaryotes, [1–3] such as cell mitosis, DNA replication, [4,5] RNA transcription, [6] macromolecule trafficking, [7] and viral genome packaging [8–14]. However, the module that initiates the ATPase action has not been clearly confirmed. Identification of the starter is of great significance for assembling artificial biological motors and for elucidating their operating mechanisms. ATPases are a class of enzymes that catalyze the decomposition of adenosine triphosphate (ATP) into adenosine diphosphate (ADP) and a free phosphate ion. They serve as the main energy source for mechanical work of biomotors, just like the engine for a car. Many ATPases adopt a multisubunit ring-shaped structure to generate force via coordinated action of multiple modules, such as the P-loop, the lid, the Walker A domain, the Walker B domain, the sensor I and II, and the arginine finger. For those ring-shaped ATPases found in biomotors, conversion from chemical to mechanical energy usually takes place in a sequential manner among the subunits, coupled with conformational changes of the oligomer. While it is commonly believed that ATP binding triggers such conversion, how ATPases start the sequential process has not been explicitly investigated. The arginine finger (R-finger), Walker A (WA), and Walker B (WB) residues are three commonly conserved motifs in ATPases responsible for ATP binding and hydrolysis [15–18]. For ring-shaped ATPases such as the AAA+ family, R-fingers are found at the inter-subunit interface regulating ATPase activities [19,20]. For instance, various mutants were assayed for ATPase activity and the trans-acting Arg139 was identified to be the R-finger of TerL ATPase [21]. The gene product 16 (gp16) of bacteriophage phi29 has long been employed to study the sequential process in ring-shaped ATPases. Based on the sequence and structural analysis, the R-finger of gp16 was determined to be Arg146, after the α4 helix. WA and WB residues are also located near the R-finger with a sequence of GXXGXGKS/T and hhhhDE, respectively. In order to translocate the viral genome double-stranded DNA (dsDNA) into the preformed capsid, the multimeric gp16 ATPases have to undergo concerted steps: ATP binding, substrate binding, ATP hydrolysis, and finally conformational changes leading to the movement of the substrate. Even without knowing the exact sequence of these steps, we can reasonably assume that the halt of the very initial step will impair all the subsequent functional steps. A recent study using the bacteriophage phi29 DNA packaging motor as a model system to examine the steps of sequential action reports that the initiator of the biological motor is the arginine finger, rather than the generally believed ATP. Through biochemical assays, it is shown that the DOI: 10.1201/9780429203367-15

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R-finger of gp16 is indispensable for subunit dimerization, ATP binding, DNA binding, and viral assembly. The alteration or removal of one amino acid residue of the arginine finger impedes or eliminates all steps, including subunit dimerization, ATP binding, DNA binding, and virus assembly. Together with molecular modeling and simulations, it is shown that the structural motifs of gp16 mediate the subunit-subunit interaction in a trans-acting manner. It has also been found that gp16 without an active R-finger can rescue ATP hydrolysis activity of an inactive Walker A mutant by trans-complementation. Therefore, we suggest that the R-finger of gp16 facilitates the dimerization of two adjacent subunits and initiates the sequential energy conversion process, finally leading to DNA translocation. Since the arginine finger directly affects subunit dimerization, the fact that arginine finger mutation impairs the ATPase function and DNA binding indicates that the dimerization of gp16 is the prerequisite of all subsequent steps for the functioning of gp16 ATPase ring. If, in natural conditions, the ATP binding and hydrolysis steps happen before the dimerization, these processes would not need an intact arginine finger to proceed. It is expected that the dimerization via trans-action of the arginine finger and the Walker A domain is a common structural feature in the AAA+ ATPases to initiate the function of biological motors. Identification of this motor initiation mechanism will also benefit the assembly of artificial biomotors in vitro and our understanding of the complex mechanism behind biopolymer translocation.

REFERENCES

1. Guo P, Schwartz C, Haak J, Zhao Z. 2013. Discovery of a new motion mechanism of biomotors similar to the earth revolving around the sun without rotation. Virology 446:133–143. 2. Guo P, Zhao Z, Haak J, Wang S, Wu D, Meng B, Weitao T. 2014. Common mechanisms of DNA translocation motors in bacteria and viruses using one-way revolution mechanism without rotation. Biotechnology Advances 32:853–872. 3. Guo P, Noji H, Yengo CM, Zhao Z, Grainge I. 2016. Biological nanomotors with a revolution, linear, or rotation motion mechanism. Microbiology and Molecular Biology Reviews 80:161–186. 4. Borowiec JA, Dean FB, Bullock PA, Hurwitz J. 1990. Binding and unwinding—how T antigen engages the SV40 origin of DNA replication. Cell 60:181–184. 5. Chakraverty RK, Hickson ID. 1999. Defending genome integrity during DNA replication: a proposed role for RecQ family helicases. Bioessays 21:286–294. 6. Gogol EP, Seifried SE, von Hippel PH. 1991. Structure and assembly of the Escherichia coli transcription termination factor rho and its interactions with RNA I. Cryoelectron microscopic studies. Journal of Molecular Biology 221:1127–1138. 7. Miyata H, Nishiyama S, Akashi K-i, Kinosita K. 1999. Protrusive growth from giant liposomes driven by actin polymerization. Proceedings of the National Academy of Sciences 96:2048–2053. 8. Guo P, Peterson C, Anderson D. 1987. Prohead and DNA-gp3-dependent ATPase activity of the DNA packaging protein gp16 of bacteriophage φ29. Journal of Molecular Biology 197:229–236. 9. Wolfe A, Phipps K, Weitao T. 2014. Viral and cellular SOS-regulated motor proteins: dsDNA translocation mechanisms with divergent functions. Cell & Biosciences 4:31. 10. Lebedev AA, Krause MH, Isidro AL, Vagin AA, Orlova EV, Turner J, Dodson EJ, Tavares P, Antson AA. 2007. Structural framework for DNA translocation via the viral portal protein. The EMBO Journal 26:1984–1994. 11. Kondabagil K, Draper B, Rao VB. 2012. Adenine recognition is a key checkpoint in the energy release mechanism of phage T4 DNA packaging motor. Journal of Molecular Biology 415:329–342. 12. Harjes E, Kitamura A, Zhao W, Morais MC, Jardine PJ, Grimes S, Matsuo H. 2012. Structure of the RNA claw of the DNA packaging motor of bacteriophage ϕ29. Nucleic Acids Research 40:9953–9963. 13. Dixit AB, Ray K, Black LW. 2012. Compression of the DNA substrate by a viral packaging motor is supported by removal of intercalating dye during translocation. Proceedings of the National Academy of Sciences 109:20419–20424. 14. Catalano C. 2000. The terminase enzyme from bacteriophage lambda: a DNA-packaging machine. Cellular and Molecular Life Sciences CMLS 57:128–148. 15. Boyer PD. 1993. The binding change mechanism for ATP synthase—some probabilities and possibilities. Biochimica et Biophysica Acta (BBA)-Bioenergetics 1140:215–250.

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16. Boyer PD. 1997. The ATP synthase—a splendid molecular machine. Annual Review of Biochemistry 66:717–749. 17. Abrahams JP, Leslie AG, Lutter R, Walker JE. 1994. Structure at 2.8 A resolution of F1-ATPase from bovine heart mitochondria. Nature 370:621. 18. Ogura T, Whiteheart SW, Wilkinson AJ. 2004. Conserved arginine residues implicated in ATP hydrolysis, nucleotide-sensing, and inter-subunit interactions in AAA and AAA+ ATPases. Journal of Structural Biology 146:106–112. 19. Zhao Z, Zhang H, Shu D, Montemagno C, Ding B, Li J, Guo P. 2017. Construction of asymmetrical hexameric biomimetic motors with continuous single‐directional motion by sequential coordination. Small 13:1601600. 20. Zhao Z, De-Donatis GM, Schwartz C, Fang H, Li J, Guo P. 2016. Arginine finger regulates sequential action of asymmetrical hexameric ATPase in dsDNA translocation motor. Molecular and Cellular Biology 36:2514–2523. 21. Hilbert BJ, Hayes JA, Stone NP, Duffy CM, Sankaran B, Kelch BA. 2015. Structure and mechanism of the ATPase that powers viral genome packaging. Proceedings of the National Academy of Sciences 112:E3792–E3799.

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Three-Step Channel Conformational Changes Common to DNA Translocases of Bacterial Viruses T3, T4, SPP1, and phi29 Shaoying Wang The Ohio State University University of Kentucky

Zhouxiang Ji The Ohio State University

Erfu Yan University of Kentucky

Farzin Haque and Peixuan Guo The Ohio State University University of Kentucky

CONTENTS 15.1 Introduction........................................................................................................................... 160 15.2 Materials and Methods.......................................................................................................... 161 15.2.1 Materials and Reagents.............................................................................................. 161 15.2.2 Expression and Purification of phi29, SPP1, T3, and T4 Portals............................... 162 15.2.3 Preparation of Lipid Vesicles Containing the phi29, SPP1, T4, and T3 Portals........ 162 15.2.4 Portal Insertion into Planar Lipid Bilayer................................................................. 162 15.2.5 Electrophysiological Measurements.......................................................................... 162 15.3 Results.................................................................................................................................... 163 15.3.1  Cloning, Expression, and Purification of the Portals of phi29, SPP1, T4, and T3...... 163 15.3.2 Insertion of Portal Channels into Lipid Membrane for Determining Channel Size Using Conductance Measurements���������������������������������������������������� 163 15.3.3 Three-Step Gating of phi29, SPP1, T4, and T3 Portal Channels.............................. 165 15.4 Discussion.............................................................................................................................. 165 15.5 Conclusions............................................................................................................................ 167 Author Contributions...................................................................................................................... 167 Acknowledgments........................................................................................................................... 167 References....................................................................................................................................... 168

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INTRODUCTION

DNA translocation motors are ubiquitous in all living systems (Guo et al., 2014; Noji and Yoshida, 2001; Grainge, 2010; Grainge, 2008; Aksyuk et al., 2009; Chung et al., 2011). During replication, the genome of double-stranded DNA (dsDNA) viruses is packaged into a preformed protein shell, referred as prohead, to a density similar to that of crystalline DNA (Earnshaw and Casjens, 1980). This process requires a powerful ATP-utilizing packaging motor to accomplish the task. In many viruses, the motor consists of DNA translocase that hydrolyzes ATP as it packages DNA and an auxiliary subunit that is usually a protein oligomer, but in bacteriophage phi29, an RNA component is required for genomic DNA packaging. The packaging RNA (pRNA) is located between the capsid shell and the ATPase motor ring (Guo et al., 1998; Hendrix, 1998). In many dsDNA bacteriophages and herpes viruses, the motor docks onto a structure called the portal. Structural studies have shown that portals from herpes virus and different tailed bacteriophages, such as phi29, SPP1, T4, and T3, share a similar cone-shaped dodecamer structure (Figure 15.1), even though their primary sequences do not display homology. The portal plays a critical role in genome packaging and ejection. During assembly, it serves as a docking point for the motor ATPase and a conduit for dsDNA transport. After DNA packaging, the portal then serves as a binding site for tail components to complete virion assembly. When bacteriophages initiate infection, DNA is ejected through the coaxial channel of the portal and tail channel into the host cell. In bacteriophage SPP1, the portal protein undergoes a concerted structure conformational change during its interaction with DNA (Chaban et al., 2015). Recent results from both membrane-embedded phi29 portal protein and DNA packaging assays in vitro demonstrated that dsDNA moves in only one direction from the narrow part to the wide end of the portal, which was referred as “oneway traffic”. More recent studies have shown that the DNA translocation of bacteriophage phi29

FIGURE 15.1 Structures of phi29, SPP1, and T4 and T3 portal channels. Top view, side view, and single subunit of phi29 (a), SPP1 (b), and T4 (c) portal protein. Phi29 gp10 PDB: 1FOU; SPP1 gp6 PDB: 2JES; T4 gp20 PDB: 3JA7.

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uses a “revolving through one-way valve” mechanism by hexameric motor (Fang et al., 2012; Jing et al., 2010b; Guo, 2014) rather than the pentameric rotational model (Morais et al., 2008; Liu et al., 2014) to package DNA, and T4 has been proposed to utilize a “compression” mechanism (Ray et al., 2010; Dixit et al., 2012; Petrov and Harvey, 2008). Since the channels act like a one-way valve, an obvious question arises: how dsDNA is ejected during infection if the channel is a one-way inward valve? Earlier studies have demonstrated that the portal exercises conformation changes during DNA packaging and ejection processes. For example, one of the studies showed that phi29 portal conformation change was induced by DNA, pRNA, or divalent metal ions assayed by circular dichroism and quenching of intrinsic tryptophan fluorescence (Urbaneja et al., 1994; Tolley and Stonehouse, 2008). Cryo-EM has also revealed conformational changes of free in vitro portal and the portal in the infectious virion (Tang et al., 2008). However, none of these studies to date have shown the conformation changes at the single-molecule level. Nanopore-based single-molecule detection has attracted immense attention from many disciplines with versatile applications recently, for example, the detection of small molecules including chemicals, nucleotides, drugs, enantiomers, as well as larger polymers such as PEG, polypeptides, RNA, and DNA (Haque et al., 2013b; Branton et al., 2008; Venkatesan and Bashir, 2011; Healy, 2007; Majd et al., 2010; Kasianowicz et al., 2008; Howorka and Siwy, 2009; Reiner et al., 2012). One novel application is the insertion of the portal from bacteriophage phi29 into an artificial membrane to serve as a unique nanopore (Wendell et al., 2009) for single-molecule detection of chemicals (Haque et al., 2012), and diagnosis of disease (Wang et al., 2013). This technology is a label-free and requires only trace amounts of analyte without the need for sample preparation or amplification. Detection can be carried out with ultra-high sensitivity within a very short time in the presence of large amount of non-specific material. We previously inserted the engineered phi29 gp10 portal into a lipid bilayer, where the portal channel shows highly robust electrophysiological properties that withstand a wide range of solution conditions, including pH 2–12 and 0.1–3M salt concentrations (Wendell et al., 2009; Jing et al., 2010a). Insertion of the portal channel into a lipid membrane results in distinguishable step size increases in current, and the channel exhibits equal conductance under both positive and negative voltage (Wendell et al., 2009; Jing et al., 2010a). By introducing appropriate probes in the interior, or at either end of the channel, single chemical or single antibody molecule can be detected at ultra-low concentrations based on the distinctive current signatures (Wang et al., 2013; Haque et al., 2012). The channel also allows dsDNA translocation (Wendell et al., 2009; Jing et al., 2010b; Haque et al., 2015; Geng et al., 2013; Geng et al., 2011; Fang et al., 2012; Haque and Guo, 2011) and is able to discriminate ssDNA and ssRNA with appropriate modification (Geng et al., 2013). Furthermore, the phi29 portal channel displays voltage-induced channel gating properties comparable to the oneway traffic property for dsDNA translocation during DNA packaging (Geng et al., 2011; Jing et al., 2010b; Fang et al., 2012). Here, we report the finding of conformational changes in the channel that are common to bacteriophages T3, T4, SPP1, and phi29. These observations support the idea that the one-way inbound channel is transformed into an outbound channel during DNA ejection.

15.2

MATERIALS AND METHODS

15.2.1 MateRials and Reagents The phospholipid, 1,2-diphytanoyl-sn glycerol-3-phosphocholine (DPhPC) (Avanti Polar Lipids), n-decane (Fisher), and chloroform (TEDIA) were used as instructed by the vendor. All other reagents were from Sigma, if not specified. Lipid A: 5% (wt/vol) DPhPC in hexane. Lipid B: 20% (wt/vol) DPhPC in decane. His binding buffer: 15% glycerol, 0.5 M NaCl, 5 mM imidazole, 10 mM ATP, 50 mM Tris-Cl, pH 8.0). His washing buffer: 15% glycerol, 500 mM NaCl, 50 mM imidazole, 10 mM ATP, 50 mM Tris-HCl, pH 8.0. His elution buffer: 15% glycerol, 500 mM NaCl, 500 mM imidazole, 50 mM ATP, 50 mM Tris-Cl, pH 8.0.

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15.2.2 expRession and puRiFiCation oF pHi29, spp1, t3, and t4 poRtals The expression and purification of phi29 portal followed the procedure reported previously (Haque et al., 2013a; Wendell et al., 2009). Gene 6 encoding SPP1 portal protein gp6 and gene 8 encoding T3 portal protein gp8 were synthesized (Genescript) and then cloned separately into the pET3c vector between the NdeI and BamHI sites. A 6×His-tag was inserted at the C-terminus for purification. Plasmids harboring genes 6 or gene 8 were transformed separately into Escherichia coli BL21(DE3), and a single colony was cultured in 10 mL Luria-Bertani (LB) medium overnight at 37°C. The culture was transferred to 1 L of fresh LB medium, and 0.5 mM IPTG was added to induce protein expression after the OD600 reached 0.5–0.6. After 3 hours, cells were collected by centrifugation at 6,000×rpm for 15 minutes, and the pellet was resuspended in His binding buffer. Bacteria were lysed by passing through a French press, and the clear supernatant was collected after centrifugation at 12,000×rpm for 20 minutes and then loaded onto a HisBind® Resin Column. SPP1 or T3 portal protein was eluted from the His•Bind® Resin Column with His elution buffer after several rounds of washing. T4 gene 20 gene, encoding the portal protein gp20, was amplified from the T4 genome and cloned into pET3c at the NdeI site and BamHI site (Keyclone). A 6×His-tag was introduced at the C-terminus for purification. Due to its hydrophobicity, T4 portal easily aggregates. Protein expression and purification were therefore modified (Quinten and Kuhn, 2012). Plasmid pET3c harboring gene 20 was transformed into E. coli HMS174(DE3), and a single colony was cultured in 10 mL LB medium overnight at 37°C. The culture was transferred to 1 L of fresh LB medium and cultured until OD600 reached 0.5–0.6. IPTG (0.5 mM final concentration) was then added to induce T4 portal protein expression. The culture was transferred to 18°C, and incubation continued overnight. Cells were harvested by centrifugation at 6,000×rpm for 15 minutes and resuspended in His binding buffer. Cells were lysed by passing through a French press. The cell pellet was collected after centrifugation at 12,000×rpm for 20 minutes and resuspended in His binding buffer containing 1% N-lauroylsarcosine for 20 minutes. The supernatant was collected after centrifugation at 12,000 × rpm for 1 hour and loaded to HisBind® Resin Column and eluted after several rounds of washing. The purity of all final protein products was verified by 10% SDS-PAGE gel.

15.2.3 pRepaRation oF lipid vesiCles Containing tHe pHi29, spp1, t4, and t3 poRtals All portal/liposome complexes were prepared following the procedure used previously (Wendell et al., 2009; Haque et al., 2013a). Briefly, 0.5 mL of 1 mg/mL DPhPC in chloroform was added to a round-bottomed flask, and the chloroform was evaporated under vacuum using a rotary evaporator (Buchi). The dried lipid film was rehydrated with 0.5 mL of portal protein solution containing 250 mM sucrose. Unilamellar lipid vesicles were obtained by extruding the lipid solution through a 400-nm polycarbonate membrane filter (Avanti Polar Lipids).

15.2.4

poRtal inseRtion into planaR lipid bilayeR

Insertion into a lipid bilayer with portal reconstituted liposomes has been reported previously (Haque et al., 2013a; Wendell et al., 2009). Briefly, a thin Teflon partition with an aperture of 200 μm was used to separate the bilayer lipid membrane (BLM) cell into cis- and trans-compartments. The aperture was pre-painted with 5% (wt/vol) DPhPC in hexane solution. The cis- and trans-chambers were filled with conducting buffer, 1 M KCl, 5 mM HEPES, pH 7.8. Then, 20% (wt/vol) DPhPC in decane solution was used to form a lipid bilayer. After confirming the formation of the lipid bilayer, the portal/liposome complexes were added to the cis-chamber to fuse with the planar lipid bilayer to form the membrane-embedded nanopore.

15.2.5 eleCtRopHysiologiCal MeasuReMents The stochastic nanopore-sensing technique is based on the principle of the classical Coulter Counter or the “resistive-pulse” routine (Coulter, 1953). The portal is located in an electrochemical chamber,

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which is separated into two compartments filled with conducting buffers. Under an applied voltage, ions passing through the portal channel will generate current in pico-Ampere (pA) scale (Haque et al., 2013b). When a charged molecule passing through the channel, it will generate transient current blockade caused by the exclusion of ions from the pore. Various parameters, such as the event dwell time, current amplitude, and unique electrical signature of the current blockages, can be used either individually or in combination for detection. A pair of Ag/AgCl electrodes inserted into both compartments was used to measure the current traces across the BLM. The current trace was recorded using an Axopatch 200B patch clamp amplifier coupled with the BLM workstation (Warner Instruments) or the Axon DigiData 1440A analogdigital converter (Axon Instruments). All voltages reported were those of the trans-compartment. Data were low-band-pass-filtered at a frequency of 5 kHz or 1 kHz and acquired at a sampling frequency of 2 kHz. PClamp 9.1 software (Axon Instruments) was used to collect the data, and Origin Pro 8.0 was used for data analysis.

15.3 15.3.1

RESULTS Cloning, expRession, and puRiFiCation oF tHe poRtals oF pHi29, spp1, t4, and t3

Following the strategy previously used for the purification of phi29 portal (Haque et al., 2013a; Wendell et al., 2009), a 6  ×  His tag was inserted at C-terminus of the SPP1, T4, and T3 portal channels to facilitate purification. A 6  ×  glycine linker was introduced between the portal and His-tag to provide flexibility. Both phi29 and SPP1 portals are soluble in the cytoplasm of E. coli. The T4 portal showed a strong tendency to aggregate due to its hydrophobic nature. Therefore, 1% N-lauroylsarcosine surfactant was added to the purification buffer to solubilize the protein (Quinten and Kuhn, 2012). After purification to homogeneity, proteins were analyzed by 10% SDS-PAGE. The single protein subunit of the phi29, SPP1, T4, and T3 portals corresponded to their predicted molecular weights of 36 kDa, 56 kDa, 60 kDa, and 59 kDa, respectively. In bacteriophage phi29, the portal is composed of 12 protein subunits that assemble into a ring to form a central channel (Sun et al., 2010). According to the crystal structure (Guasch et al., 2002), the dimensions of phi29 portal are: 13.8 nm and 6.6 nm in diameter at the wide and narrow ends, respectively. In SPP1, the channel assembles as 13-mer and the narrowest part is 2.77 nm (Figure 15.1) (Lebedev et al., 2007; Lhuillier et al., 2009). An early EM study revealed that the T4 portal exists as a dodecameric ring, with contour dimensions of 14 nm long and 7 nm wide, and about ~3 nm in diameter in the interior of the channel (Driedonks et al., 1981); a more recent structure gives the external diameter as 9 nm at the upper end, 12 nm in the middle, and 8 nm at the lower end. The internal diameter of central channel is about 4.4 nm at the upper end and about 3 nm in the narrowest part (Enoki et al., 2015). However, no crystal structure of the portal is currently available. Several studies have revealed that the T3 portal protein is a mixed population of 12 and 13 subunits. The percentages of these two oligomer states vary by culture, indicating that assembly of the portal depends on the expression conditions and other factors (Valpuesta et al., 1992; Carazo et al., 1986; Valpuesta et al., 2000). EM studies revealed that the three-dimensional structure of the 12-mer T3 portal is as follows: 14.9 nm at external diameter; 8.5 nm in height; and an average 3.7 nm at internal open channel (Agirrezabala et al., 2005; Valpuesta et al., 1992; Valpuesta et al., 2000).

15.3.2 inseRtion oF poRtal CHannels into lipid MeMbRane FoR deteRMining CHannel size using ConduCtanCe MeasuReMents To incorporate phi29, SPP1, T4 and T3 portal proteins into planar lipid bilayers, we adopted the twostep procedure described previously (Wendell et al., 2009): reconstitution of portal in liposomes, followed by the fusion of proteoliposomes with planar lipid bilayer to form the membrane-embedded

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portal protein. Experiments were carried out using the same buffer 1 M KCl, 5 mM HEPES, pH 7.8, under 50 mV applied potential. The current jump for each channel insertion was measured at a fixed voltage to determine the channel sizes of the phi29, SPP1, T3, and T4 portal channels (Figure 15.2a– d). Each current jump represents the insertion of one channel into the lipid bilayer. Since fusion of the portal protein/liposome with the planar lipid bilayer is a random event, the time between independent insertion events varies. Figure 15.2a–d provides representative results for the portals of the four phages. The channel conductance (derived from the ratio of measured current jump to the applied voltage) of phi29, SPP1, and T4 was determined to be 4.52 ± 0.33 nS, 4.10 ± 0.22 nS, and 3.03 ± 0.37 nS, respectively (Figure 15.2e–g). T3 conductance distribution appeared as two peaks: 2.65 ± 0.31 nS and 3.90 ± 0.38 nS (Figure 15.2h), which likely correspond to the 12-mer and 13-mer portal channels. Under a scanning voltage (‒50 mV to +50 mV; 2.2 mV/s), the phi29, SPP1, T4, and T3 portal channels all display a linear current-voltage (I–V) relationship without voltage gating (Figure 15.2i–l). When 100 mV was applied, the phi29, T4, and T3 portal channels remained stable, again without displaying voltage gating, but the SPP1 portal channel started gating at this voltage (data not shown). In addition, the SPP1 portal channel also has a stronger tendency to gate under a negative voltage than under a positive gradient (data not shown).

FIGURE 15.2 Representative current traces showing the insertion of phi29 (a), SPP1 (b), T4 (c), and T3 (d) portal channels into planar lipid membrane. Applied voltage: +50 mV. Histogram showing the conductance distribution of phi29 (e), SPP1 (f), T4 (g), and T3 (h) portal channels. Applied voltage: +50 mV. Current-voltage trace under a ramping potential (‒50 mV to +50 mV; 2.2 mV/s) for phi29 (single channel) (i), SPPI (two channels) (j), T4 (one channel) (k), and T3 (three channels) (l) portals. Conducting buffer: 1 M KCl, 5 mM HEPES, pH 7.8.

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15.3.3 tHRee-step gating oF pHi29, spp1, t4, and t3 poRtal CHannels When a higher voltage (>100 mV) was applied, three distinct steps of conformational change in the channel are observed in all four portal channels. Conformational changes in the channel are reflected in a reduction in electrical current of 33%, 66%, and 99% for the first, second, and third step, respectively (Figure 15.3). Three discrete steps of gating of the phi29, SPP1, T4, and T3 portal channels are found under an applied positive voltage of 150 mV, 150 mV, 170 mV, and 150 mV, respectively (Figure 15.3a–d). Similar phenomena were observed under negative voltages of ‒125 mV, ‒100 mV, ‒175 mV, and ‒125 mV (Figure 15.3e–h). These are the minimum voltages required for gating the channels.

15.4

DISCUSSION

The polymorphism of portal complex assembled from overexpressed gene of bacteriophages has been reported for many years. Although it is believed that the T4 and SPP1 portals exist as dodecamers in biologically active state (Cardarelli et al., 2010; Doan and Dokland, 2007; Guasch et al., 2002; Cingolani et al., 2002; Badasso et al., 2000; Cuervo and Carrascosa, 2011; Xiao et al., 2009; Rodriguez-Casado et al., 2001), the stoichiometry of the portal in

FIGURE 15.3 Three-step gating associated with conformational changes in phi29 (a), SPP1 (b), T4 (c), and T3 (d) portal channels under positive trans-membrane voltages. Three-step gating associated with conformational changes in phi29 (e), SPP1 (f), T4 (g), and T3 (h) portal channels under negative transmembrane voltage.

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different bacteriophages has been reported to vary from 11-mer to 14-mer in vitro following expression and assembly (Cingolani et al., 2002; Dube et al., 1993; Kocsis et al., 1995; Trus et al., 2004; Valpuesta et al., 2000; Rao and Feiss, 2008; Camacho et al., 2003; Tsuprun et al., 1994). Several studies revealed that T3 portal structure is a mixed population of 12 and 13 subunits (Valpuesta et al., 1992). The percentages of these two oligomer states vary by preparation, indicating that the portal protein assembly depends on the expression conditions and other factors (Valpuesta et al., 1992; Carazo et al., 1986; Valpuesta et al., 2000). The diverse distribution of conductance for phi29, SPP1, T4, and T3 portals might represent various oligomer states in these portal complexes. This is reflected in the displaying of two major peaks in T3 conductance distribution (Figure 15.2h). Further studies have revealed that specific SPP1 portal segments, such as helix α6 of the tunnel loop and the crown region, may be responsible for different oligomeric states (Lebedev et al., 2007). Studies on T4 portal by Cryo-EM revealed that the ratio of the 11-mers, 12-mers, and 13-mers shifted after loop deletion (Sun et al., 2015). It has been proved that all portal channels of dsDNA bacteriophages display a left-handed channel wall configuration to facilitate one-way traffic of dsDNA into procapsid by a revolution mechanism without rotation (Jing et al., 2010b; Zhao et al., 2013; Schwartz et al., 2013; De-Donatis et al., 2014). The one-way valve mechanism is consistent with the finding of genome gating in SPP1, albeit the gating mechanism proposed by these authors is based on the analysis of the channel structure after the completion of DNA entry instead of during translocation (Ziolo et al., 2005). The finding of the “pushing through a one-way valve” mechanism (Fang et al., 2012; Zhang et al., 2012; Jing et al., 2010b) raises the question of how dsDNA is ejected during infection if the channel only permits dsDNA to translocate in one direction. We believe during translocation of the dsDNA, the interaction of the dsDNA with the channel wall and the procapsid component next to the portal will trigger conformational changes in the portal; therefore, the left-handed portal channel, which facilitates dsDNA advancement with one direction, will transition to a neutral or right-handed configuration in three steps to facilitate the DNA ejection after DNA packaging is complete (Geng et al., 2011; De-Donatis et al., 2014). Such conformational changes in portal proteins as proposed above for ejection of the packaged dsDNA have previously been reported (Tang et al., 2008; Geng et al., 2011; Tang et al., 2008; Guo et al., 2005). It has been shown that DNA, pRNA, or divalent metal ions could induce conformational change in phi29 portal as revealed by circular dichroism measurement and the quenching study on the intrinsic tryptophan fluorescence (Urbaneja et al., 1994; Tolley and Stonehouse, 2008). Portal gate closing has been reported in SPP1 (Orlova et al., 2003) and speculated in T4 (Sun et al., 2015). It was reported recently that the portal of bacteriophage SPP1 undergoes a concerted reorganization of the structural elements of its central channel during interaction with DNA. Structural rearrangements and gate closing were reported to associate with protein-protein and protein-DNA interaction, and a diaphragm-like mechanism for channel reduction and gate closing has been proposed (Chaban et al., 2015). It was speculated that tunnel loops within the channel serve as contact point for DNA to trigger conformational change (Orlova et al., 2003; Grimes et al., 2011). Loop mutation assay led to the speculation that these loops are involved in the retention of the packaged DNA (Grimes, Ma et al., 2011) and in the preparation for motor detachment before tail addition to complete the virion assembly (Chaban et al., 2015). Protein-protein interactions prime the channel loops to build a plug to close the channel to prevent genome leakage from the capsid. The gatekeeper system opens for viral genome exit at the beginning of infection but recloses afterward, suggesting a molecular diaphragm-like mechanism to control DNA efflux. We found that within the phi29 system, the migration of DNA through the motor channel requires a voltage in the range of 15–150 mV. If the voltage is smaller than 15 mV, dsDNA translocation is not detectable because DNA translocation needs to overcome the enthalpy and entropy costs. If the voltage is higher than 150 mV, channel gating will occur. In vivo studies have revealed a threshold voltage

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in the range of 60–90 mV that was needed for the ejection of genome from T4 phage (Labedan et al., 1980). Although the in vivo systems are different from the condition in this report, it is still valid that an energy level is required to open or close the portal channel. In this study, three steps of conformational changes for the portal of phi29, SPP1, T4, and T3 were observed. Such conformational changes would allow conversion of the left-handed portal after completion of DNA packaging towards the opposite configuration, thus facilitating DNA one-way ejection into host cells for infection. Furthermore, these gating observations may be a universal property of all portals. A diaphragm mechanism (Chaban et al., 2015) has been proposed, and our finding for the three steps might be an interpretation with the analogy of a camera lens by suggesting discrete f-stops, like f4.5, f8, f16, f32. However, the diaphragm proposal is difficult to reconcile with the data by Baker and coworkers {5972} who reported a right-handed twisting of the connector structure in comparing the free connector with the structure of the connector in the DNA-filled virion {5972}. It was reported that when treated as a rigid body, the left-handed 30° tilt in the crystal structure of phi29 connector does not fit into the Cryo-EM density maps of the DNAfilled phi29 virion, as demonstrated in a correlation coefficient as low as 0.55. The correlation coefficient was improved to 0.70 after manual adjustment, resulting in a 10° twist of the portal towards the portal axis {5972}. On the other hand, it’s difficult to adjust to fit the N-terminal external region as a rigid body into other parts of the connector density of the crystal structure of the free connector. It was discovered that the N-terminal external region underwent a significant conformational shift in the DNA-filled capsid compared with the free connector structure {5972}. It has been concluded that restructuring of the connector core subunit and angular twisting are promoted by the interactions among its structural proteins and phi29 DNA {5972}. Due to the relatively static C-terminal internal region and the dsDNA alignment with the channel wall {7964,8755,7803,7804, 7940}, a significant conformational change in the N-terminal external region can lead to a clockwise twist of the dsDNA when viewed from the C-terminus, which agrees with the reported twisting as observed by optical tweezers {8848}. It is possible that the three gating steps may also correspond to the quantized steps of partial genome ejection observed in T3 {9731} and the partial packaging intermediates observed in phi29 {242} (Serwer et al., 2014; Bjornsti et al., 1983).

15.5 CONCLUSIONS The motor channel of T3, SPP1, T4, and phi29 all display three discrete steps of voltage gating resulting from channel conformational changes, suggesting that the one-way inbound channel during the DNA packaging process is transformed into an outbound channel prepared for DNA ejection during the host infection.

AUTHOR CONTRIBUTIONS We thank Zhengyi Zhao, Daniel Binzel, and Mario Vieweger for insightful comments. PG, SW, and FH designed research; SW, JX, and EY performed research; SW, JX, and EY analyzed data; SW, PG, and FH wrote the paper.

ACKNOWLEDGMENTS The research was supported by NIH Grant R01EB012135 and funding to Guo’s Endowed Chair in Nanobiotechnology position from the William Fairish Endowment Fund. Peixuan Guo is cofounder of Kylin Therapeutics, Inc., and Biomotor and RNA Nanotechnology Development Corp. Ltd.

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Geng, J., Fang, H., Haque, F., Zhang, L., Guo, P., 2011. Three reversible and controllable discrete steps of channel gating of a viral DNA packaging motor. Biomaterials 32, 8234–8242. Geng, J., Wang, S., Fang, H., Guo, P., 2013. Channel size conversion of phi29 DNA-packaging nanomotor for discrimination of single- and double-stranded nucleic acids. ACS Nano 7, 3315–3323. Grainge, I., 2008. Sporulation: SpoIIIE is the key to cell differentiation. Current Biology 18, R871–R872. Grainge, I., 2010. FtsK--a bacterial cell division checkpoint? Molecular Microbiology 78, 1055–1057. Grimes, S., Ma, S., Gao, J., Atz, R., Jardine, P. J., 2011. Role of phi29 connector channel loops in late-stage DNA packaging. Journal of Molecular Biology 410, 50–59. Guasch, A., Pous, J., Ibarra, B., Gomis-Ruth, F. X., Valpuesta, J. M., Sousa, N., Carrascosa, J. L., Coll, M., 2002. Detailed architecture of a DNA translocating machine: the high-resolution structure of the bacteriophage phi29 connector particle. Journal of Molecular Biology 315, 663–676. Guo, P., 2014. Biophysical studies reveal new evidence for one-way revolution mechanism of bacteriophage phi29 DNA packaging motor. Biophysical Journal 106, 1837–1838. Guo, P., Grainge, I., Zhao, Z., Vieweger, M., 2014. Two classes of nucleic acid translocation motors: rotation and revolution without rotation. Cell & Bioscience 4, 54. Guo, P., Zhang, C., Chen, C., Trottier, M., Garver, K., 1998. Inter-RNA interaction of phage phi29 pRNA to form a hexameric complex for viral DNA transportation. Molecular Cell 2, 149–155. Guo, Y., Blocker, F., Xiao, F., Guo, P., 2005. Construction and 3-D computer modeling of connector arrays with tetragonal to decagonal transition induced by pRNA of phi29 DNA-packaging motor. Journal of Nanoscience and Nanotechnology 5, 856–863. Haque, F., Geng, J., Montemagno, C., Guo, P., 2013a. Incorporation of viral DNA packaging motor channel in lipid bilayers for real-time, single-molecule sensing of chemicals and double-stranded DNA. Nature Protocols 8, 373–392. Haque, F., Guo, P., 2011. Membrane-embedded channel of bacteriophage phi29 DNA-packaging motor for translocation and sensing of double-stranded DNA. In: S. M. Iqbal, R. Bashir (Eds.), Nanopores, Sensing and Fundamental Biological Interactions. Springer, pp. 77–106. Haque, F., Li, J., Wu, H.-C., Liang, X.-J., Guo, P., 2013b. Solid-state and biological nanopore for real-time sensing of single chemical and sequencing of DNA. Nano Today 8, 56–74. Haque, F., Lunn, J., Fang, H., Smithrud, D., Guo, P., 2012. Real-time sensing and discrimination of single chemicals using the channel of phi29 DNA packaging nanomotor. ACS Nano 6, 3251–3261. Haque, F., Wang, S., Stites, C., Chen, L., Wang, C., Guo, P., 2015. Single pore translocation of folded, double-stranded, and tetra-stranded DNA through channel of bacteriophage phi29 DNA packaging motor. Biomaterials 53, 744–752. Healy, K., 2007. Nanopore-based single-molecule DNA analysis. Nanomedicine 2, 459–481. Hendrix, R. W., 1998. Bacteriophage DNA packaging: RNA gears in a DNA transport machine (minireview). Cell 94, 147–150. Howorka, S., Siwy, Z., 2009. Nanopore analytics: sensing of single molecules. Chemical Society Reviews 38, 2360–2384. Jing, P., Haque, F., Shu, D., Montemagno, C., Guo, P., 2010b. One-way traffic of a viral motor channel for double-stranded DNA translocation. Nano Letters 10, 3620–3627. Jing, P., Haque, F., Vonderheide, A., Montemagno, C., Guo, P., 2010a. Robust properties of membraneembedded connector channel of bacterial virus phi29 DNA packaging motor. Molecular BioSystems 6, 1844–1852. Kasianowicz, J. J., Robertson, J. W., Chan, E. R., Reiner, J. E., Stanford, V. M., 2008. Nanoscopic porous sensors. Annual Review of Analytical Chemistry 1, 737–766. Kocsis, E., Cerritelli, M. E., Trus, B. L., Cheng, N., Steven, A. C., 1995. Improved methods for determination of rotational symmetries in macromolecules. Ultramicroscopy 60, 219–228. Labedan, B., Heller, K. B., Jasaitis, A. A., Wilson, T. H., Goldberg, E. B., 1980. A membrane potential threshold for phage T4 DNA injection. Biochemical and Biophysical Research Communications 93, 625–630. Lebedev, A. A., Krause, M. H., Isidro, A. L., Vagin, A. A., Orlova, E. V., Turner, J., Dodson, E. J., Tavares, P., Antson, A. A., 2007. Structural framework for DNA translocation via the viral portal protein. EMBO Journal 26, 1984–1994. Lhuillier, S., Gallopin, M., Gilquin, B., Brasiles, S., Lancelot, N., Letellier, G., Gilles, M., Dethan, G., Orlova, E. V., Couprie, J., Tavares, P., Zinn-Justin, S., 2009. Structure of bacteriophage SPP1 head-to-tail connection reveals mechanism for viral DNA gating. Proceedings of the National Academy of Sciences of the United States of America 106, 8507–8512.

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Liu, S., Chistol, G., Hetherington, C. L., Tafoya, S., Aathavan, K., Schnitzbauer, J., Grimes, S., Jardine, P. J., Bustamante, C., 2014. A viral packaging motor varies its DNA rotation and step size to preserve subunit coordination as the capsid fills. Cell 157, 702–713. Majd, S., Yusko, E. C., Billeh, Y. N., Macrae, M. X., Yang, J., Mayer, M., 2010. Applications of biological pores in nanomedicine, sensing, and nanoelectronics. Current Opinion in Biotechnology 21, 439–476. Morais, M. C., Koti, J. S., Bowman, V. D., Reyes-Aldrete, E., Anderson D, Rossman, M. G., 2008. Defining molecular and domain boundaries in the bacteriophage phi29 DNA packaging motor. Structure 16, 1267–1274. Noji, H. and Yoshida, M., 2001. The rotary machine in the cell, ATP synthase. Journal of Biological Chemistry 276(3), 1665–1668. Orlova, E. V., Gowen, B., Droge, A., Stiege, A., Weise, F., Lurz, R., van, H. M., Tavares, P., 2003. Structure of a viral DNA gatekeeper at 10 A resolution by cryo-electron microscopy. EMBO Journal 22, 1255–1262. Petrov, A. S., Harvey, S. C., 2008. Packaging double-helical DNA into viral capsids: structures, forces, and energetics. Biophysical Journal 95, 497–502. Quinten, T. A., Kuhn, A., 2012. Membrane interaction of the portal protein gp20 of bacteriophage T4. Journal of Virology 86, 11107–11114. Rao, V. B., Feiss, M., 2008. The bacteriophage DNA packaging motor. Annual Review of Genetics 42, 647–681. Ray, K., Sabanayagam, C. R., Lakowicz, J. R., Black, L. W., 2010. DNA crunching by a viral packaging motor: compression of a procapsid-portal stalled Y-DNA substrate. Virology 398, 224–232. Reiner, J. E., Balijepalli, A., Robertson, J. W., Campbell, J., Suehle, J., Kasianowicz, J. J., 2012. Disease detection and management via single nanopore-based sensors. Chemical Reviews 112, 6431–6451. Rodriguez-Casado, A., Moore, S. D., Prevelige, P. E., Thomas, G. J., 2001. Structure of bacteriophage P22 portal protein in relation to assembly: investigation by Raman spectroscopy. Biochemistry 40, 13583–13591. Schwartz, C., De Donatis, G. M., Zhang, H., Fang, H., Guo, P., 2013. Revolution rather than rotation of AAA+ hexameric phi29 nanomotor for viral dsDNA packaging without coiling. Virology 443, 28–39. Serwer, P., Wright, E. T., Liu, Z., Jiang, W., 2014. Length quantization of DNA partially expelled from heads of a bacteriophage T3 mutant. Virology 456–457, 157–170. Sun, L., Zhang, X., Gao, S., Rao, P. A., Padilla-Sanchez, V., Chen, Z., Sun, S., Xiang, Y., Subramaniam, S., Rao, V. B., Rossmann, M. G., 2015. Cryo-EM structure of the bacteriophage T4 portal protein assembly at near-atomic resolution. Nature Communications 6, 7548. Sun, S., Rao, V. B., Rossmann, M. G., 2010. Genome packaging in viruses. Current Opinion in Structural Biology 20, 114–120. Tang, J. H., Olson, N., Jardine, P. J., Girimes, S., Anderson, D. L., Baker, T. S., 2008. DNA poised for release in bacteriophage phi29. Structure 16, 935–943. Tolley, A. C., Stonehouse, N. J. 2008. Conformational changes in the connector protein complex of the bacteriophage phi29 DNA packaging motor. Computational and Mathematical Methods in Medicine 9, 327–337. Trus, B. L., Cheng, N., Newcomb, W. W., Homa, F. L., Brown, J. C., Steven, A. C., 2004. Structure and polymorphism of the UL6 portal protein of herpes simplex virus type 1. Journal of Virology 78, 12668–12671. Tsuprun, V., Anderson, D., Egelman, E. H., 1994. The Bacteriophage phi29 head-tail connector shows 13-fold symmetry in both hexagonally packed arrays and as single particles. Biophysics Journal 66, 2139–2150. Urbaneja, M. A., Rivqas, S., Carrascosa, J. L., Valpuesta, J. M., 1994. An intrinsic-tryptophan-fluorescence study of phage phi29 connector/nucleic acid interactions. European Journal of Biochemistry 225, 747–753. Valpuesta, J. M., Fujisawa, H., Marco, S., Carazo, J. M., Carrascosa, J., 1992. Three-dimensional structure of T3 connector purified from overexpressing bacteria. Journal of Molecular Biology 224, 103–112. Valpuesta, J. M., Sousa, N., Barthelemy, I., Fernandez, J. J., Fujisawa, H., Ibarra, B., Carrascosa, J. L., 2000. Structural analysis of the bacteriophage T3 head-to-tail connector. Journal of Structural Biology 131, 146–155. Venkatesan, B. M., Bashir, R., 2011. Nanopore sensors for nucleic acid analysis. Nature Nanotechnology 6, 615–624. Wang, S., Haque, F., Rychahou, P. G., Evers, B. M., Guo, P., 2013. Engineered nanopore of phi29 DNApackaging motor for real-time detection of single colon cancer specific antibody in serum. ACS Nano 7, 9814–9822. Wendell, D., Jing, P., Geng, J., Subramaniam, V., Lee, T. J., Montemagno, C., Guo, P., 2009. Translocation of double-stranded DNA through membrane-adapted phi29 motor protein nanopores. Nature Nanotechnology 4, 765–772.

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16

Sequence Dependence of Reversible CENP-A Nucleosome Translocation Micah P. Stumme-Diers, Thomas Stormberg, and Yuri L. Lyubchenko University of Nebraska Medical Center

CONTENTS 16.1 Introduction .......................................................................................................................... 173 16.2 Results and Discussion ......................................................................................................... 174 16.3 Materials and Methods ......................................................................................................... 176 Acknowledgements ........................................................................................................................ 178 References ...................................................................................................................................... 178

16.1

INTRODUCTION

The generation of two genetically identical daughter cells from a parent cell is achieved through mitosis when sister chromatids are divided evenly between the newly formed cells. Critical to this process is a specialized locus possessed by all chromosomes called the centromere, which serves as the point of spindle fiber attachment via the kinetochore when distributing the chromatids to the daughter cells. Without a functioning centromere, chromosome segregation becomes aberrant resulting in aneuploidy, a hallmark of cancer [1]. The characteristic feature defining centromere nucleosomes is the presence of centromere protein A (CENP-A), which replaces histone H3 in nucleosomes at this chromatin region [2,3]. Unlike canonical nucleosomes containing histone H3 that wraps DNA in ~1.75 turns around the histone core, in nucleosomes containing CENP-A the histone core wraps only 1.5 DNA turns [4–6]. We recently proposed a model that describes the structural consequences of nucleosomes and their arrays that may arise when DNA makes only 1.5 turns around the histone core and how it can help to distinguish the centromere from bulk chromatin [7]. It was further hypothesized that the dynamic pathways followed by CENP-A nucleosomes are unique from those with H3, which likely play a role in centromere maintenance and function. In line with this hypothesis, single-molecule biophysics methods capable of probing the transient states of nucleosome dynamics have found CENP-A nucleosomes to be highly dynamic and capable of spontaneous disassembly [8–11]. In a recent study, we performed direct imaging of CENP-A nucleosomes by high-speed time-lapse atomic force microscopy (HS-AFM) and revealed several dynamic pathways unique to CENP-A nucleosomes that were not observed for H3 nucleosomes [11]. The spontaneous unwrapping of DNA from the CENP-A core was found to be accompanied by the dynamic formation of loops of DNA at the size of one wrap of DNA. Loops were also found to mediate the reversible translocation of a CENP-A core along the DNA substrate, eventually resulting in the transfer of the nucleosome core to a new, previously unoccupied DNA substrate. An intriguing property of the translocation process is the stepwise manner in which it progresses, suggesting a possible sequence dependence of the process. To address this, we performed further analysis of this translocation process, and the results are presented here. DOI: 10.1201/9780429203367-17

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FIGURE 16.1 Reversible translocation of CENP-A nucleosome core along DNA substrate. Snapshots of the forward and reverse translocation of the histone core are shown above the curve. The reverse movement of the core brings it back to about the same position on the DNA substrate as it began. The white arrow in image one indicates the arm for which the contour length measurements were made. The distance measured from the end of the arm indicated above to the center of the core particle for every frame of the video. White scale bar indicates 20 nm. Black circles represent raw data, and the black line a moving median.

16.2

RESULTS AND DISCUSSION

The translocation process of the CENP-A histone core was measured as the distance from one end of the DNA to the center of the core. As shown in Figure 16.1, the core begins on the 601 positioning sequence close to the center of the DNA substrate (frame 1) where it remains until moving to the end of the DNA substrate while stopping at various points along the way. Quantitative analysis of these data is shown in Figure 16.1 as a graph in which time-dependent translocation of the nucleosome core in both directions is shown. The translocation is not a perfectly monotonous process; rather, there are positions at which the core stalls. This stepwise translocation also takes place when the core moves in the reverse direction, back to the center of the substrate until resettling in the same position as it began (frames 6–9). We plot data shown in Figure 16.1 as a histogram in which residence time at specific locations is calculated from the number events at selected areas (Figure 16.2a). This plot reveals two peaks positioned at ~150 and ~68 bp from the end of the substrate at which the nucleosome almost stalls. When determining preferential occupancy, we found that the nucleosome is nearly four times as likely to stall on an area of elevated GC content during the translocation to the end of the DNA. A similar plot for the reversal translocation reveals two peaks at positions ~65 and ~160 bp, along with a peak at ~position 115 bp from the end of the substrate (Figure 16.2b). We calculated the GC content variation of this DNA segment and plotted these graphs in Figure 16.2a and b as red lines. The GC content was calculated over 20-bp windows. The comparison of the two plots reveals that the nucleosome core stalls at DNA segments with elevated GC content (Figure 16.2a). The preferential occupancy, while still significant, is less pronounced during the translocation to the center, as the nucleosome is just over two times as likely to occupy an area of elevated GC

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FIGURE 16.2 Histograms of the CENP-A core position during translocation. (a) Translocation from the middle to end of the substrate is seen to stall twice as indicated by the two peaks positioned at ~150 and ~68 bp from the substrate end. (b) Translocation from the end to the middle of the substrate stalls three times as evident by the three peaks of the histogram. In both a and b, GC% at each distance from the end is represented as a black curve overlaying on the plot, and the direction of translocation is indicated by the arrow above the plots.

content. The end-to-middle translocation follows a similar trend, with the stalled movement at ~65 and ~160 bp happening at similar regions of high GC content as translocation in the other direction (Figure 16.2b). Overall, the nucleosome is shown to preferentially occupy segments of elevated GC content approximately three times as often as segments with lower GC content. In stalls of 10 or more frames, the histone core occupied an area elevated GC content 73% of the time. From this analysis, we suggest a moderate positive correlation between GC content and nucleosome core occupancy (r = 0.4862, p = < 0.001). We analyzed an additional extended nucleosome dynamic event observed in our paper [11], where a nucleosome was shown to thread DNA into a loop before rewrapping after a short translocation. We performed the analysis with the goal to determine if the relationship between GC content and nucleosome occupancy existed in a different type of the nucleosome translocation event. The separate arm lengths of the complex were measured on a frame-by-frame basis to determine which DNA segments were being newly occupied by the nucleosome during the looping and unlooping process. The arm lengths over time are plotted in Figure 16.3. In this plot, we can see that the looping process

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FIGURE 16.3 Arm length measurements of looping event. Snapshots of the looping and unlooping of the histone core are shown above the plot. White scale bar indicates 20 nm. The top arrow in image one indicates arm 1, and the bottom arrow indicates arm 2. The plot indicates the length of each arm and the sum of arms during each frame of the event. Arm 1 undergoes a significant change, while arm 2 stays relatively stable.

predominantly takes place on arm 1 of the nucleosome process. We then plotted how the GC content of DNA taken in by the nucleosome core changed over time. Figure 16.4a shows that arm 1 of the complex begins at an unfavorably low GC content level. The DNA continues to loop through the core until reaching an elevated level of GC content and closing. Figure 16.4b shows that the GC content fluctuates throughout the looping process, but the arm stays relatively stable and closes at the same GC content level that it began with. Overall, this process increases the GC content from 45% before the loop is formed to 64% when the loop is closed. Our finding on the nucleosome dynamics depending on the GC content is in line with a recently reported relationship between elevated GC content and increased nucleosome occupancy [12]. We note that our data represent the observations of only two separate translocation events. Further studies on the sequence-dependent nucleosome dynamics are needed to address the sequencedependent nucleosome translocation phenomenon found in these studies. Specifically, studies on human alpha-satellite DNA and other centromere-specific DNA would provide both detail on the sequence dependence of nucleosome translocation and other dynamic properties of nucleosomes, including the looping of DNA from the core.

16.3

MATERIALS AND METHODS

These methods are an abbreviated version of those previously reported for these experiments [11]. Nucleosomes containing a CENP-A octamer core were assembled using salt gradient dialysis with a 423-bp substrate containing a 147-bp 601 positioning sequence flanked by plasmid DNA 154 and 122 bp in length. The stability and quality of nucleosome assemblies were checked by discontinuous SDS-PAGE (Sodium Dodecyl Sulfate-PolyAcryl Amide Electrophoresis) gel with 6% stacking and 15% separating components and using AFM imaging in ambient conditions. From the AFM imaging, it was determined that CENP-A nucleosomes from the assembly wrap 122.3 ± 15 bp, which agrees well with the 121 bp expected. For HS-AFM imaging, nucleosome samples were diluted to ~1–2 nM in a buffer containing 10 mM HEPES (pH 7.5) and 4 mM MgCl2 followed immediately by deposition on freshly cleaved mica surfaces functionalized with a 167 μM

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177

FIGURE 16.4 Comparison of complex arm length to GC content occupied by the histone core over time. (a) The length of arm 1 undergoes a drastic change throughout the looping process. The GC content rises from an unfavorable 33% before the looping begins to 71% after the loop closes. (b) The length of arm 2 stays relatively stable throughout the looping process. The GC content both begins and ends at 52%. The cumulative GC% rises from 45% to 64%. In both a and b, GC% at each distance from the end is represented as a black curve overlaying on the plot and the arm length is represented as a black curve.

1-(3-aminopropyl)-silatrane (APS) solution. After a 2-minute incubation, the sample is exchanged for imaging buffer (same as dilution) and imaged using HS-AFM (RIBM) using an Olympus Micro Cantilever (BL-AC10DS-A2) that was EBD-treated. Typical images were acquired at a scan rate of 0.2-0.4 sec/frame at a 200 × 200 nm size. Measurements of histone core sliding and looping were done frame by frame using FemtoScan Online analysis software. Since one of the substrate arms drifts outside of the frame while capturing the sliding movie, only one arm was measured. This was achieved by measuring the contour length of the DNA from its end to the center of the histone core. The GC content along the DNA strand was determined by calculating what percentage of each 20-bp rolling segment contained GC. For example, if a 20-bp segment contains 3 G, 7 C, 5 A, and 5 T, the %GC for this segment would be 50%. If a segment contains 10 G, 5 C, 2A, and 3 T, the %GC for the segment would be 75%.

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ACKNOWLEDGEMENTS This work was supported by NSF Grant MCB 1515346 and MCB 2123637, NIH Grants GM096039 and GM118006, all to YLL.

REFERENCES 1. Gordon DJ, Resio B, Pellman D (2012) Causes and consequences of aneuploidy in cancer. Nat Rev Genet 13 (3):189–203 2. Black BE, Jansen LET, Maddox PS, Foltz DR, Desai AB, Shah JV, Cleveland DW (2007) centromere identity maintained by nucleosomes assembled with histone H3 containing the CENP-A targeting domain. Mol Cell 25 (2):309–322. doi:10.1016/j.molcel.2006.12.018. 3. McKinley KL, Cheeseman IM (2016) The molecular basis for centromere identity and function. Nat Rev Mol Cell Biol 17 (1):16–29. doi:10.1038/nrm..2015.5. 4. Tachiwana H, Kagawa W, Kurumizaka H (2012) Comparison between the CENP-A and histone H3 structures in nucleosomes. Nucleus 3 (1):6–11 5. Tachiwana H, Kagawa W, Shiga T, Osakabe A, Miya Y, Saito K, Hayashi-Takanaka Y, Oda T, Sato M, Park S-Y, Kimura H, Kurumizaka H (2011) Crystal structure of the human centromeric nucleosome containing CENP-A. Nature 476 (7359):232–235. http://www.nature.com/nature/journal/v476/n7359/ abs/nature10258.html - supplementary-information 6. Yoda K, Ando S, Morishita S, Houmura K, Hashimoto K, Takeyasu K, Okazaki T (2000) Human centromere protein A (CENP-A) can replace histone H3 in nucleosome reconstitution in vitro. Proc Nat Acad Sci 97 (13):7266–7271. doi:10.1073/pnas..130189697. 7. Lyubchenko YL (2014) Centromere chromatin: a loose grip on the nucleosome? Nat Struct Mol Biol 21 (1):8-8. doi:10.1038/nsmb.2745. 8. Lyubchenko YL (2014) Nanoscale nucleosome dynamics assessed with time-lapse AFM. Biophys Rev 6 (2):181–190. doi:10.1007/s12551-013-0121-3. 9. Shlyakhtenko LS, Lushnikov AY, Lyubchenko YL (2009) Dynamics of nucleosomes revealed by timelapse atomic force microscopy. Biochemistry 48 (33):7842–7848. 10. Menshikova I, Menshikov E, Filenko N, Lyubchenko YL (2011) Nucleosomes structure and dynamics: effect of CHAPS. Int J Biochem Mol Biol 2:129–137. 11. Stumme-Diers MP, Banerjee S, Hashemi M, Sun Z, Lyubchenko YL (2017) Nanoscale dynamics of centromere nucleosomes and the critical roles of CENP-A. Nucleic Acids Res. doi:10.1093/nar/gkx933. 12. Kaplan N, Moore IK, Fondufe-Mittendorf Y, Gossett AJ, Tillo D, Field Y, LeProust EM, Hughes TR, Lieb JD, Widom J, Segal E (2009) The DNA-encoded nucleosome organization of a eukaryotic genome. Nature 458 (7236):362–366. doi:10.1038/nature07667

17

Same Function from Different Structures among pac Site Bacteriophage (TerS) Terminase Small Subunits Lindsay W. Black and Krishanu Ray University of Maryland School of Medicine

CONTENTS References ...................................................................................................................................... 182 Many years of experimental study has supported the basic similarity of phage and viral DNA packaging motor mechanisms. The similar high-resolution structures of proheads, portals, and large terminase subunits (TerLs) support this view. The terminase small subunit (TerS) is essential in vitro and in vivo for packaging of circular or concatemeric DNA (i.e., DNA with no or few ends), the latter being the in vivo substrate that is subject to complex regulation of packaging initiation among pac site phages. In pac site phages, the TerS is required to initiate packaging at specific pac sites on concatemeric DNA by “handing off” the DNA to the large nuclease-containing terminase subunit (TerL) for packaging initiation cutting followed by ATP-driven DNA translocation into the prohead by TerL (Figure 17.1a). In vitro, in contrast, among phages such as T4, TerL acting alone can package linear DNA of any sequence and size with high efficiency; evidently, recognition of a DNA end by TerL obviates the need for TerS to initiate and continue DNA translocation into the prohead. Substantial evidence supports a double TerS protein ring/double DNA pac site “synapsis model” as a mechanism able to assess DNA concatemer maturation sufficient for the initiation of DNA packaging and prohead filling by TerL [1] (Figure 17.1). In fact, there is both strong genetic and biochemical evidence for this proposed mechanism for the initiation of phage T4 packaging: (i) purified TerS displays enhanced binding to a GC-rich sequence at the 3’ end of its gene that was identified as a pac site [2]; (ii) this sequence confers enhanced transduction of phage and plasmid DNAs containing it in vivo by a transducing derivative of T4 [3]; (iii) under selection for increased synthesis of TerL, the TerS protein is shown to be required for gene amplifications requiring recombination between the homologous gene 16 and 19 pac sequences (see Figure 17.1b) [4]; such sequencespecific amplifications require a phage internal Alt protein knockout to allow substantially more DNA to be packaged into the phage particle, apparently by increasing head volume, thus allowing multiple copies of the 16–19 region to be viable within the mature chromosome [5,6]; (iv) a mature DNA restriction fragment containing a gene 16 pac site can be found in phage particles [3]; however, site-directed mutagenesis of the gene 16 amplification pac site that eliminates gene 16–19 amplifications is not lethal to the phage, suggesting backup pac sites (such as the homologous 19 pac site (Figure 17.1b) or other mechanisms [7]; and finally (v) mass spectrometry and scanning transmission electron microscopy mass determination show single 11mer and double 22mer TerS T4 rings (Figure 17.2); these are proposed to be aplanar, lock washer-like single- and double-ring structures that unstack to yield side-by-side rings [2]. The single and double rings purified from an untagged protein expression vector are stable by mass spectrometry and have been shown to contain only DOI: 10.1201/9780429203367-18

179

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Biomotors and their Nanobiotechnology Applications (a)

capsid shell (gp23)

“Terminase small subunit (TerS) twin ring synapsis model"

dsDNA

portal vertex (gp20)

(c) multimeric terminaseATPase large subunit gp17 (TerL) ring

pac

packaging

(b) concatemer

pac 16 16 pac

4 kb

16

pac 170 kb

recombination

pac

19 small terminase ring (gp16) 16-19 gene amplification

16pac - GAGGCTCAAGAAGCTCGTGAGAAG 19pac - GAAGCTCATGATGCTCGTCAGAAG

--Twin ring nucleosome-like structure: pairing of chromosome homologs -measures DNA replication for packaging -Two pac duplex DNAs paired: strand swapped Holliday junction signal

FIGURE 17.1 Conserved bacteriophage two subunit terminase pac site prohead packaging; (a) large terminase-ATPase subunit TerL (T4 gp17) alone packages linear DNAs in vitro; (b) small terminase subunit TerS (T4 gp16) is required in vivo for concatemer (or circular DNA) packaging; a “synapsis model” proposes DNA concatemer maturation is assessed by TerS apposing two homologous pac site DNAs. Sequence-specific terminase gene amplifications requiring TerS (gp16) result from apposition of the two homologous pac sites (16 and 19) shown under genetic selection for increased TerL synthesis; normally, 16-16 pac apposition would initiate packaging; and (c) a two ring two dsDNA TerS Holliday junction strand swap signal is proposed to initiate handoff from TerS to TerL for DNA cutting and packaging.

protein [8]. The two rings thus are not held together with DNA as hypothesized for another proposed two-ring gp16 structure, although without direct experimental support [9]. Unlike crystal structures of His-tagged gp16 that by mass form 11mers and 12mers, the purified untagged rings are by mass only 11mers (not 12mers) and 22mers. Of course, in vivo the TerS protein is expected to interact with DNA, possibly as shown in Figure 17.1c. An unexpected difference between TerL and TerS is that while the large terminase protein has long been known to be a signature homology feature among diverse phages [10]. For current structure-based TerL informatics [by S. Hardies see Figure 2 in Ref. [11]], the small terminase genes and gene product crystal structures display marked differences. Moreover, major controversies abound as to how these crystal structures relate to TerS DNA binding in various pac site phages. Thus, whether the pac DNA-binding site is at the N-terminus of TerS (where there is generally found to be a DNA-binding motif) or at its C-terminus, and how this relates to number of monomers per crystal ring structure (8, 9, 10, 11, and 12 per single ring have been reported among different phages), as well as the very different TerS monomer structures [for a gallery of the single-ring crystal structures. see Figure 17.2 [12], and for the side-by-side double T4 TerS protein only rings, see Figure 17.2 [2]], and whether DNA moves through a central ring channel and/or wraps around the ring is debated [9,12–16]. Even more importantly, the relevance of the rings to function remains problematic since the single planar rings of variable numbers of monomers from different pac site phages have been produced at high protein expression levels or observed under crystallization conditions. Interestingly,

Same Function from Different Structures among pac Site Phage T4 TerS 11mers and 22mers

181

Mass in phage T4 TerS 11mers and 22mers

FIGURE 17.2 Shown in the top panels are phage T4 TerS 11mer and 22mer protein-only rings; the relative amounts of the 11mers and 22mers are established by scanning transmission electron microscopy molecular weight determination [2]; the double-headed arrow scale bar is 6.7 nm. In the bottom panel are shown the highly variable TerS protein crystal structures from a number of pac site phages, although phage ƛ is not a pac site phage because its TerS operates by a strict DNA sequence-specific TerS + TerL mechanism rather than a TerS plus pac site-only mechanism.

phage SPP1 pac cleavage is relatively precise without a DNA sequence specificity requirement, supporting the importance of DNA bending for targeting pac site cutting by TerS [17]. The T4 terminase small subunit TerS protein remains active when modified at both the N-terminal and C-terminal ends. In fact, gp16-mCherry and gp16-GFP fusion proteins are highly fluorescent and readily purified as N-terminal His-tag proteins. Moreover, both these His-tag expression vectorsynthesized proteins are active in vivo as judged by their complementation with gene 16 amber mutants. Full activity is also observed when TerS-mCherry and TerS-GFP fusion protein-encoding genes replace the normal terS gene by recombination into the T4 genome. Full TerS activity is retained despite the marked increase in its molecular weight (18–45 kDa). The existence of fully functional N- and C-terminal gp16 fusions is contrary to proposals that the DNA moves through the small-ring central channel [14] and an argument against the proposal that DNA binding is to the C-terminal end of the ring monomer [12]. At least in phage T4 bulky C-terminal fusion, additions to gp16 would be expected to affect DNA binding and DNA translocation through a narrow-diameter TerS channel. FRET shows that a ring form of TerS is apparently produced in vivo at low expression levels by co-infection using the two recombinant phage genomes; moreover, in contrast to wild type, when purified, a temperature-sensitive (ts) mutant form of gp16 forms rings at 20°C (permissive) but not 42°C (non-permissive), strongly suggesting that TerS rings are produced and required for function in vivo. In fact, if it is assumed that a DNA-wrapped single ring or double ring as shown in Figure 17.1c is the active TerS structure, the great variability in numbers of monomers and structures in different phage TerS rings likely obscures quite similar functions and DNA structures. Our TerS gene fusion results support the consensus view that DNA initially binds the N-terminal portion

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of the small subunit, where the DNA-binding motif is located [2], and then wraps around the smallsubunit ring without passing through the central channel. A DNA wrapping structure in which DNA bending and Holiday junction formation may play as important a role as DNA sequence-specific binding fits well to the double protein-double DNA ring structure. Taken among all the pac site phages, and emphasizing the T4 TerS double ring, the evidence in favor of a nucleosome-like TerS ring discussed above for a double protein ring – double DNA pac structure is strong. It remains to be firmly established whether the proposed critical signal for packaging initiation is the Holliday junction structure formed between swapped strands in the four-stranded pac site paired structure as shown in Figure 17.1c. This proposed mechanism is favored by the pac sequence-specific, TerS-required specific gene amplifications that can arise because of recombination between apposed homologous gene 16 and gene 19 pac sequences under strong genetic selection for enhanced TerL synthesis, as shown in Figure 17.1b and as referenced above. Alternatively, the formation of the double DNA double protein ring may be sufficient, without Holiday junction strand swapping, to act as the signal for the TerL initiation cutting and subsequent packaging. In vitro demonstration of Holiday junction formation of apposed 16–19 DNA sequence rings by purified TerS could further support the proposed recombinational mechanism. Research reported in this publication was supported by the National Institute of General Medical Sciences of the National Institutes of Health under Award Numbers (R01GM118766 to LWB and R01 GM117836 to KR). The content is solely the responsibility of the authors and does not necessarily represent the official views of the National Institutes of Health.

REFERENCES 1. Black LW. 1995. DNA packaging and cutting by phage terminases: control in phage T4 by a synaptic mechanism. Bioessays 17:1025–1030. 2. Lin H, Simon MN, Black LW. 1997. Purification and characterization of the small subunit of phage T4 terminase, gp16, required for DNA packaging. J Biol Chem 272:3495–3501. 3. Lin H, Black LW. 1998. DNA requirements in vivo for phage T4 packaging. Virology 242:118–127. 4. Wu CH, Lin H, Black LW. 1995. Bacteriophage T4 gene 17 amplification mutants: evidence for initiation by the T4 terminase subunit gp16. J Mol Biol 247:523–528. 5. Wu DG, Black LW. 1987. Gene amplification mechanism for the hyperproduction of T4 bacteriophage gene 17 and 18 proteins. J Mol Biol 195:769–783. 6. Wu DG, Wu CH, Black LW. 1991. Reiterated gene amplifications at specific short homology sequences in phage T4 produce Hp17 mutants. J Mol Biol 218:705–721. 7. Wu CH, Black LW. 1995. Mutational analysis of the sequence-specific recombination box for amplification of gene 17 of bacteriophage T4. J Mol Biol 247:604–617. 8. van Duijn E. 2010. Current limitations in native mass spectrometry based structural biology. J Am Soc Mass Spectrom 21:971–978. 9. Sun S, Gao S, Kondabagil K, Xiang Y, Rossmann MG, Rao VB. 2012. Structure and function of the small terminase component of the DNA packaging machine in T4-like bacteriophages. Proc Natl Acad Sci U S A 109:817–822. 10. Black LW. 1989. DNA packaging in dsDNA bacteriophages. Annu Rev Microbiol 43:267–292. 11. Serwer P, Jiang W. 2012. Dualities in the analysis of phage DNA packaging motors. Bacteriophage 2:239–255. 12. Roy A, Bhardwaj A, Datta P, Lander GC, Cingolani G. 2012. Small terminase couples viral DNA binding to genome-packaging ATPase activity. Structure 20:1403–1413. 13. Buttner CR, Chechik M, Ortiz-Lombardia M, Smits C, Ebong IO, Chechik V, Jeschke G, Dykeman E, Benini S, Robinson CV, Alonso JC, Antson AA. 2012. Structural basis for DNA recognition and loading into a viral packaging motor. Proc Natl Acad Sci U S A 109:811–816. 14. Zhao H, Finch CJ, Sequeira RD, Johnson BA, Johnson JE, Casjens SR, Tang L. 2010. Crystal structure of the DNA-recognition component of the bacterial virus Sf6 genome-packaging machine. Proc Natl Acad Sci U S A 107:1971–1976. 15. Roy A, Cingolani G. 2012. Structure of p22 headful packaging nuclease. J Biol Chem 287:28196–28205. 16. Teschke CM. 2012. Themes and variations of viral small terminase proteins. Structure 20:1291–1292. 17. Djacem K, Tavares P, Oliveira L. 2017. Bacteriophage SPP1 pac cleavage: a precise cut without sequence specificity requirement. J Mol Biol 429:1381–1395.

18

Kinetic Study of the Fidelity of DNA Replication with Higher-Order Terminal Effects Yao-Gen Shu

Wenzhou Institute, University of Chinese Academy of Sciences

CONTENTS 18.1 Introduction .......................................................................................................................... 183 18.2 Basic Theory of Steady-State Copolymerization Kinetics ................................................... 184 18.2.1 Bernoullian Model: Zero-Order Terminal Effects ................................................... 185 18.2.2 Terminal Model: First-Order Terminal Effects ........................................................ 187 18.2.3 Penultimate Model: Second-Order Terminal Effects ............................................... 189 18.2.4 Higher-Order Terminal Models ................................................................................ 191 18.3 DNA Replication: A Binary Copolymerization in Two Dimensions ................................... 192 18.3.1 Basic Theory of Steady-State Kinetics of the Exonuclease Proofreading Model .... 194 18.3.1.1 First-Order Proofreading Model ................................................................ 194 18.3.1.2 Second-Order Proofreading Model ........................................................... 196 18.3.2 The Fidelity of DNA Replication.............................................................................. 197 18.3.2.1 The Infinite-State Markov Chain Method for Exonuclease Proofreading ..... 198 18.3.2.2 Approximation of φ under Bio-Relevant Conditions .................................200 18.4 Case Study: T7 DNA Polymerase ........................................................................................202 18.5 Discussion and Conclusion ................................................................................................... 203 Acknowledgments.......................................................................................................................... 203 References ...................................................................................................................................... 203

18.1

INTRODUCTION

Since the Watson-Crick (WC) base-pairing rules of double-strand DNA were discovered [1], template-directed DNA replication has become a critical research subject to understand genetic variations and evolution. It is now widely acknowledged that WC pairings (match: A-T and G-C) play a dominating role in the replication process so as to maintain the genome stability, while the non-WC pairings (mismatch) occur with very low probability (about 10 −4 –10 −10, dependent on species). This is not due to the difference between the free energy of match and mismatch in the double-helical DNA: in fact, this free energy difference is only about 2 ∼ 4kBT, which cannot account for such low error rates (high fidelity) if estimated by Boltzmann factor. As pointed out by J. Hopfield [2] and J. Ninio [3], the low error rates may originate from the huge difference of replication kinetics between match and mismatch, which is realized by high-fidelity DNA polymerases (DNAp, a molecular motor that catalyzes the template-directed DNA synthesis) [4,5]. While experiments have revealed for a long time that the DNAp fidelity is determined by template-dependent synthesis kinetics, related theoretical problems were not solved, e.g., how does the template sequence influence the kinetics and the statistical features of the nascent chain? how to calculate the positional fidelity (reciprocal of the error rate at each template position) or the overall DOI: 10.1201/9780429203367-19

183

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fidelity correctly, if all the involved kinetic parameters are experimentally measured? Systematic theoretical studies on these issues appeared quite recently. So far, there are two categories of models. One assumes that the kinetic parameters of all match (or mismatch) are of similar orders of magnitude (i.e., without considering the template sequence explicitly) and thus describes the synthesis approximately as a match/mismatch binary copolymerization process. This simplification has long been used in biochemistry to analyze experimental data or to build theoretical models (e.g., see the historical literatures [2,3] or more recent publications like [6–9]). However, models of this category have not been systematically studied until recently, especially for cases in which the rates of unit addition or deletion at the end of the growing chain depend on the preceding one or more units. Such higher-order neighbor effects may be significant if there is a mismatch at the terminal region of the growing chain to destabilize the terminus and hence affect the unit addition or deletion. These effects can now be properly handled under the steady-state assumptions, and the overall replication fidelity and growth velocity can be calculated numerically or analytically [10–12]. In these treatments, the steady-state assumption plays a central role to simplify the copolymerization process as a homogenous Markov chain. This is, however, not appropriate for real cases in which the template DNA sequence is highly inhomogenous and the kinetic parameters of match/mismatch are highly sequence dependent. These template-sequence specificities have not received much attention until very recently. All the 16 types of base pairs can now be explicitly considered in the theoretical models, and the nearest neighbor effects can be handled properly [13–17]. By these models, one can compute the local fidelity or velocity for any position along the template (i.e., the fidelity or velocity profile), rather than the estimates of the overall fidelity or velocity. However, there are still many open questions. For instance, the fidelity profile for a given template sequence can be computed by the iteration algorithm, which goes through the entire sequence cyclically for convergence [17]. This seems to imply that the fidelity for any position may depend on the entire sequence, which is doubtful, for it is hard to conceive that replication mutations at different positions have long-range correlations rather than randomly distributed as widely believed. To what a range do the positional quantities depend on the surrounding template sequence? Do the correlations in the template sequence (if any) have any influence on the fidelity profile? In this chapter, we review the kinetic treatment to the copolymerization process as a homogenous Markov chain with the steady-state assumption. We first introduce a basic theory of steady-state copolymerization kinetics in one dimension [11]. Then, we treat the DNA replication as a binary copolymerization in two dimensions due to proofreading [12]. Finally, we will apply our method to treat the T7 DNAp as a first-order case [18].

18.2

BASIC THEORY OF STEADY-STATE COPOLYMERIZATION KINETICS

Understanding the kinetics of copolymerization and thus controlling the copolymer sequence statistics (e.g., copolymer composition, sequence distribution) are the key subjects in the study of copolymer, since the sequence statistics significantly affect the chemical and physical properties of the copolymers [19]. Therefore, it became an important issue to theoretically model the copolymerization kinetics and estimate the rate constants of all the involved polymerization reactions. This has drawn a lot of attention both experimentally and theoretically since the 1940s. In order to study the kinetics, the experiments are usually conducted at low conversion conditions to maintain the copolymerization process at steady state (i.e., the monomer concentrations in the environment are almost unchanged during the process), which actually much simplifies the modeling and analysis of experiment data. Based on the steady-state assumption, different theoretical models have been suggested for different systems. Early works assumed the so-called terminal effects, i.e., the last monomer unit at the growing end of the copolymer influences the chain growth and thus the copolymer composition. Several terminal models were developed in the 1940s and successfully applied to experiments [20–23]. Besides the assumptions of terminal effect, these early models also assumed that the

Kinetic Study of the Fidelity of DNA Replication

185

copolymerization reactions are irreversible, which ensures the corresponding kinetic equations to be solved analytically. These two assumptions were shown insufficient to explain later experimental results, which leads to the development of two other categories of models. The first category was proposed to account for the so-called penultimate effect, i.e., the next-tolast (penultimate) monomer unit at the growing end can have a substantial influence on the copolymerization kinetics (e.g., [24,25]). The original penultimate model was suggested by Merz et al. [26], and then was revised and developed (for a review, see Ref. [27]). Besides the terminal (also called as first-order terminal in this chapter) and penultimate (the second-order terminal) effects, higher-order terminal effects are also possible (e.g., antepenultimate effect [28]). But such cases have not been systematically investigated. The second category was proposed to account for depropagation effect, which brings substantial mathematical difficulty to the studies of copolymerization kinetics. Depropagation was noticed very early in the 1960s. It originated from the thermodynamic argument, i.e., all the reaction pathways are essentially reversible and depropagation may become significant at some elevated temperature. A few copolymerization systems do exhibit depropagation, Which shows substantial impacts on the copolymerization kinetics and copolymer composition (e.g., [29–31]). Such temperature effects can only be described by reversible models. However, depropagation always leads to hierarchically coupled and unclosed kinetic equations, which are hard to be solved analytically (as will be clear in later subsections). Because of this mathematical difficulty, it was until 1987 that the first systematic treatment of the first-order terminal models with depropagation was given by Kruger et al. [32]. Kruger’s approach was based on the key assumption that the copolymer sequence can be described as a first-order Markov chain (Eq. (12) in Ref. [32]). By using this assumption, Kruger et al. succeeded in reducing the original kinetic equations into closed steady-state equations. However, the validity of this assumption has not been proven rigorously or verified numerically. Moreover, how to generalize Kruger’s approach to higher-order terminal models was unclear. So far as we know, the only attempt to extend Kruger logic to penultimate models with depropagation has been made by Li et al. [33,34]. In their works, however, the first-order Markov chain assumption, which is valid only for terminal models, was inappropriately employed. This makes their penultimate model mathematically self-inconsistent (a detailed discussion will be given in Section 18.2.3). By far, there are no well-established penultimate models with depropagation available in the literature. Recently, the study on steady-state copolymerization also attracted attention from physicists who were interested to visualize the nonequilibrium copolymerization as information-generating process. In Ref. [35–37], the zero-order copolymerization model with depropagation (named as Bernoullian model in Section 18.2.1) was introduced, without giving the derivation of the steadystate equations, to discuss some interesting issues (e.g., fidelity of DNA replication). In Ref. [10], the first-order terminal model was discussed, similar to Kruger, under the assumption of the firstorder Markov chain. These works also put an emphasis on the thermodynamics of steady-state copolymerization and gave very general and interesting relations between the copolymer sequence entropy and the thermodynamic entropy production of the copolymerization system. However, there still lacks a systematic investigation on the steady-state kinetics and thermodynamics of any-order terminal model with depropagation. In this section, we will generalize Kruger’s Markov chain assumption of the copolymer sequence distribution and suggest a unified mathematical approach to solve the steady-state kinetic equations of any-order terminal model with depropagation.

18.2.1

beRnoullian Model: zeRo-oRdeR teRMinal eFFeCts

As the simplest case of copolymerization, Bernoullian model (i.e., zero-order terminal model) assumes that the propagation and depropagation of monomers are independent of the terminal monomer unit. Although it is not a good model for real copolymerization systems, it can serve as a starting

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FIGURE 18.1 A growing copolymer in one dimension.

point of our discussion. Below, we investigate a two-component (A, B) system. Generalization to more complex cases (e.g., multicomponent systems) will be given in later sections. Denoting the propagation rate constants as k A0 or k B0 , and depropagation rate constants as k A , k B , we have the reaction scheme below:

k0

k0

kA

kB

A B   → ~ A⋅, ~ ⋅ + B ←   →~ B ~ ⋅ + A←  

Imaging a single growing copolymer as shown in Figure 18.1, ∼ represents the reactive end (i.e., the growing end, being either A or B), and the occurrence probability of A·or B at the terminal is denoted as PA and PB, respectively. We define k A ≡ k A0 [ A], k B ≡ k B0 [ B], [A], [B] are monomer concentrations in the environment, which are constants during the steady-state copolymerization. Supposing at some moment the copolymer contains NA monomer A and NB monomer B, the total number of monomers N = NA + NB. They all increase with time during copolymerization, and the corresponding kinetic equations are N A ≡ J A = k A − k A PA

N B ≡ J B = k B − k B PB

(18.1)

N ≡ J tot = J A + J B JA, JB are, respectively, the overall incorporation rates of A and B. In steady-state copolymerization, d ( N A N ) dt = d ( N B N ) dt = 0, or equivalently, N A N = N A N = J A J tot and N B N = N B N = J B J tot So the overall occurrence probability of A or B in the copolymer can be expressed as QA ≡ N A N = J A J tot ,QB ≡ N B N = J B J tot . Higher order of chain-end sequence distribution Pin  i1 ( im = A or B, m = 1,2,  , n ). in…i1 denotes the chain-end sequence, with i1 representing the terminal unit, and the total number of sequence in  i1 occurring in the copolymer chain N in  i1 can be similarly defined. N. in … i1 ≡ Jin … i1 = ki1 Pin … i2 − ki1 Pin … i1 . In general, we have Pin …i1 = PAin …i1 + PBin …i1, Jin …i1 = J Ain …i1 + J Bin …i1 . We also define Jin…i1* ≡ Jin…i1 A + Jin…i1B and the overall sequence distribution Qin .....i1 ≡ N in ....i1 N = Jin  i1 J tot . The kinetic equations of Pin  i1 (n ≥ 1) can be written as

P.in…i1 = Jin…i1 − J~in…i1*

(18.2)

For example,



P.A = J A = J~ A* = J BA − J AB = k A PB − k B PA + k B PAB − k A PBA

(18.3)

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Kinetic Study of the Fidelity of DNA Replication

The existence of depropagation rates k A , k B makes these equations hierarchically coupled and hard to be solved. Fortunately, for steady-state copolymerization P.in…i1 = 0 (for any n ≥ 1), we can use the following truncation method to solve these equations. In Bernoullian model, the steady-state copolymerization kinetics is determined only by PA, PB. This means that the coupled equations are redundant and can be reduced to equations of the two basic variables PA, PB. This reduction can be achieved by the following zero-order factorization n

conjecture of the chain-end sequence distribution, Pin  i1 =

∏P

im

, which leads to

m=1

Jin ...i1

 = 

n

∏ m= 2

  Pim  Ji1 , Jin ...i1* =   



n

∏ P  j im

tot

(18.4)

m =1

From the steady-state kinetic equation 0 = P.in…i1 = Jin…i1 − J~in…i1*, we can get Ji1 = J tot Pi1

(18.5)

Therefore, each of the coupled equations is reduced to the same steady-state equation of PA, PB, J A JB = PA PB

(18.6)

Combining the normalization condition PA + PB = 1, we now obtain a set of closed equations, which gives the exact solution of the original kinetic equations (these steady-state equations have been used without derivation in Ref. [36,37]). However, Ref. [37] also provides another independent method, i.e., the infinite-state Markov chain model, to give an exact calculation of JA /JB, which is proven to be identical to that obtained from Eq. (18.6). This method can be generalized to higher-order terminal model and shown to be equivalent to our approach. Details can be found in Ref. [11]). Support of the factorization conjecture comes from the Monte-Carlo simulations by using Gillespie algorithm [38,39] (here, the rate parameters are arbitrarily chosen). One can directly simulate the steady-state copolymerization from any given initial condition of PA, PB, and obtain all the sequence statistics (e.g., the chain-end sequence distribution Pi1 , Pi2i1 , ) from a number of simulations. For simplicity, we only check the factorizations:   = Pi3 Pi2 Pi1   = Pi4 Pi3 Pi2 Pi1 

Pi2i1 = Pi2 Pi1 Pi3i2i1 Pi4 i3i2i1

(18.7)

As shown in Figure 18.2a, for arbitrary choice of rate parameters k A, kB, k A , k B (the only constraint on the parameters is that they should ensure Jtot > 0, i.e., the copolymer is growing), all the equalities hold when copolymerization reaches the unique steady state (which is determined only by rate parameters and independent of the choice of initial conditions).

18.2.2

teRMinal Model: FiRst-oRdeR teRMinal eFFeCts

The so-called terminal model (i.e., the first-order terminal model), where the propagation and depropagation of monomer A and B are dependent on the identity of the terminal monomer unit, is a

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Biomotors and their Nanobiotechnology Applications

FIGURE 18.2 Simulation verification of the zero-order (a), first-order (b), and second-order (c) factorization conjectures with 2000 samples. In (a), the illustrative rate parameters k A  =  4.0, kB  =  3.0, k A  =  2.0, k B  =  1.0. In (b), the illustrative rate parameters k AA  =  6.0, k AB  =  2.0, kBA  =  4.0, kBB  =  1.0, k AA   =  7.0, k AB  = 5.0, k BA = 3.0, k BB  = 1.0 and in (c), the illustrative rate parameters k AAA = 8.0, k ABA = 4.0, kBAA = 7.0, kBBA =  2.0, k AAB  =  1.0, k ABB  =  3.0, kBBB  =  5.0, kBBB  =  6.0, k AAA = 7.0, k ABA = 3.0, k BAA   =  8.0, k BBA = 4.0, k AAB  =  5.0, k ABB = 1.0, k BAB = 6.0, k BBB  = 2.0. Here, im = A, B, m = 1,2,3,4.

much more realistic model than Bernoullian model for real copolymerization systems. The reaction pathways in terminal model are k AA k BB    → AA⋅, ~ A ⋅ + B  ~ A ⋅ + A← → ~ AB ⋅ ←  k AA

k AB

k BA k BB    → ~ BA⋅, ~ B ⋅ + B  ~ B ⋅ + A← ← → ~ BB ⋅  k BA

k BB

Defining Jin…i2i1 ≡ ki2i1 Pin…i2 − ki2i1 Pin…i2i1 J~in…i2i1* ≡ Jin…i2i1 A+ Jin…i2i1B

(18.8)

where im = A, B (m = 1,2,…, n; n ≥ 2), we can write the corresponding kinetic equations for Pin…i2i1 (n ≥ 1) as below P.in … i2i1 = Jin … i2i1 − J~in … i2i1 *

(18.9)

The basic variables here are PAA, PAB, PBA, PBB, rather than PA, PB. Following the same logic in the previous subsection, we can reduce the hierarchically coupled equations Eq. (16.9) to an equivalent set of closed equations of PAA, PAB, PBA, PBB, by using the first-order factorization conjecture n

Pin  i2i1 =

∏ m= 2

−1

 n  Pim im −1  Pim −1  , n ≥ 3  m = 3 

∏

(18.10)

Then, the steady-state kinetic equations Pin ···i2i1 = 0 (n ≥ 2) are reduced to Ji2i1 Ji1 * = Pi2i1 Pi1

(18.11)

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Kinetic Study of the Fidelity of DNA Replication

or equivalently, J Ai J Bi = PAi PBi

(18.12)

where i = A, B. P.A = J A − J~ A∗ or P.B = J B − J B∗ (they are equivalent since PA + PB = 1) leads to another steady-state equation J AB = J BA. Finally, we get four equations for four variables J AA J BA J AB J BB = , = , J AB = J BA , PAA PBA PAB PBB

(18.13)

PAA + PAB + PBA + PBB = 1 which gives the solution of the original kinetic equations. The validity of the factorization Pi3i2i1 = Pi3i2 Pi2i1 Pi2 and the steady-state equations Eq. (18.13) can be checked by Monte-Carlo simulation. Figure 18.2b shows the factorization holds when copolymerization reaches steady state. The first-order factorization conjecture Pi3i2i1 = Pi3i2 Pi2i1 Pi2 is actually equivalent to the firstorder Markov chain assumption used in Ref. [32] and Ref. [10]. Defining transition probability p ( i2 | i1 ) ≡ Pi2i1 Pi1 , p ( A | i1 ) + p ( B | i1 ) = 1, we now can rewrite the factorization conjecture as Pi3i2i1 = p ( i3 | i2 ) p ( i2 | i1 ) Pi1 . It is worth noting that we have chosen Pi2i1, rather than Pi1 and p ( i2 | i1 ), as basic variables so as to represent the steady-state equations in a much simpler and more intuitive form.

18.2.3

penultiMate Model: seCond-oRdeR teRMinal eFFeCts

The reaction pathway of penultimate model (i.e., the second-order terminal model) can be expressed as   → ~ AAB ⋅ ~ AA ⋅ + A    ← → ~ AAA⋅ ~ AA ⋅ +B ← k AAA

k AAB

k AAA

k AAB

  → ~ ABB ⋅ ~ AB ⋅ + A    ← → ~ ABA⋅ ~ AB ⋅ +B ← k ABA

k ABB

k ABA

k ABB

k BAA k BAB   → ~ BAB ⋅ ~ BA ⋅ + A    ← → ~ BAA⋅ ~ BA ⋅ +B ← k BAA

k BAB

k BBA k BBB   → ~ BBB ⋅ ~ BB ⋅ + A    ← → ~ BAA⋅ ~ BB ⋅ +B ← k BBA

k BBB

Here, the basic variables are Pi3i2i1 ( i3 , i2 , i1 = A, B ). As in previous sections, we still define Jin  i3i2i1 ≡ ki3i2i1 Pin  i3i2 − ki3i2i1 Pin i3i2i1 Jin …i2i1 ∗ ≡ Jin …i2i1 A + Jin …i2i1B

(18.14)

where im = A, B (m = 1,2,…, n;  n ≥ 3). The kinetic equation of Pin  i3i2i1 is P.in … i3i2i1 = Jin … i3i2i1 − J~in … i3i2i1 *

(18.15)

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To solve these equations, we take the following second-order factorization conjecture n

Pin  i3i2i1 =

∏P

im im −1im − 2

m= 3

−1

 n  Pim −1im − 2  , n ≥ 4   m = 4 

∏

(18.16)

The steady-state kinetic equation P.in … i3i2i1 = 0 (n ≥ 3) can thus be reduced to Ji3i2i1 Ji2i1* = Pi3i2i1 Pi2i1

(18.17)

J Ai2i1 J Bi2i 1 = PAi2i1 PBi2i1

(18.18)

or equivalently

Now we have had five independent equations (Eq. (18.18), along with the normalization condition) for the eight variables. The rest three equations come from the remaining kinetic equations of Pi2i1 P.i2i1 = Ji2i1 − J~i2i1∗ = 0

(18.19)

These four equations are not independent, due to the normalization condition ∑ Pi2i1 = 1. So any three of them can be selected to form a closed set of equations of Pi3i2i1, for instance, J AAA J BAA J AAB J BAB = = , , PAAA PBAA PAAB PBAB J ABA J BBA J ABB J BBB = , = , PABA PBBA PABB PBBB

(18.20)

J AA = J AA* , J AB = J AB* , J BB = JBB* ,

∑

Pi3i2i1 = 1

i3 ,i2 ,i1 = A ,B

We checked the validity of Pi4 i3i2i1 = Pi4 i3i2 Pi3i2i1 Pi3i2 and the steady-state equations by Monte-Carlo simulations. Figure 18.2c shows the second-order factorization holds when copolymerization reaches the steady state. In general, if mth-order factorization conjecture (m < s) is applied to the steady-state kinetic equations P.in … i1 = 0 (n = 1,2, …) of sth-order model (see the next subsection), one can always obtain an overdetermined set of equations, which is mathematically self-inconsistent. On the other hand, higher-order (m > s) factorization conjecture is redundant for the sth-order model. We therefore conclude that the sth-order model can only be appropriately described by sth-order factorization conjecture.

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Kinetic Study of the Fidelity of DNA Replication

18.2.4

HigHeR-oRdeR teRMinal Models

The logic presented in previous subsections can be directly generalized to higher-order terminal models. Below, we list the major results for sth-order terminal model, i.e., the propagation and depropagation of A/B depend on the last s monomer units of the copolymer. Here, the basic variables are Pis +1 is ...i1 (2s+1 in total). We denote the propagation rates as kis +1is ...i1 and depropagation rates as kis +1 is i1, and also Jin … i3i2i1 ≡ kis +1 … i3i2i1 Pin … i3i2 − kis +1 … i3i2i1 Pin …i3i2i1 J~in … i2i1 * ≡ Jin … i2i1 A + Jin …i2i1B

(18.21)

where im = A, B; m = 1,2,  , n; n ≥ s + 1. The sth-order factorization conjecture is n

Pin  i1 =

∏P

im im −1  im − s

m = s +1

−1

 n  Pim −1  im − s  n ≥ s + 2   m = s + 2 

∏

(18.22)

The closed steady-state equations are derived from the following 2s equations J Ais is −1  i1 J Bis is −1  i1 = PAis is −1  i1 PBis is −1  i1

(18.23)

Jis +1is i1 Jis is −1 i1 = Pis +1is i1 Pis is −1 i1

(18.24)

or equivalently,

The kinetic equations P.is … i1 = Jis … i1 − J~is … i1 * = 0 give other 2s steady-state equations, from which any 2s − 1 equations can be chosen. Combining the normalization condition ∑ Pis +1is…i1 = 1, we finally obtain a closed set of 2s + 1 equations for 2s + 1 variables. The sth-order factorization conjecture can be rewritten equivalently as sth-order Markov chain, by defining the transition probability p ( is +1 | is  i1 ) ≡ Pis +1is  i1 Pis  i1 , p ( A is  i1 ) + p ( B is  i1 ) = 1. Noting that J~is is −1 … i1* = Jis is −1 … i1 , the steady-state equations Eq. (18.24) can be transformed into Jis +1is i1 Pis +1is i1 = Pis  i1 Jis  i1

(18.25)

Since the overall sequence distribution Qis +1is  i1 Qis  i1 = Jis +1is  i1 Jis  i1 , the steady-state equations can be rewritten as Qis +1is i1 Pis +1is i1 = = p ( is +1 is  i1 ) Qis  i1 Pis  i1

(18.26)

This simply means that the overall sequence distribution and chain-end sequence distribution can be described by the same sth-order Markov chain.

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It is also worth noting that the sth-order model can reproduce (s − 1)th-order model if assuming k Ais  i1 = k Bis  i1 = kis  i1 and k Ais  i1 = k Bis  i1 = kis  i1 . By the sth-order model, we have PAis ....i2i1 J Ais ...i2i1 kis ...i2i1 PAis ...i2 − kis ...i2i1 PAis ...i2i1 = = PBis ...i2i1 J Bis ....i2i1 kis ...i2i1 PBis ...i2 − kis ...i2i1 PBis ...i2i1

(18.27)

PAis i2i1 PAis i2 = PBis i2i1 PBis i2

(18.28)

PAis i2i1 PBis i2i1 Pis  i2i1 = = PAis i2 PBis i2 Pis  i2

(18.29)

which yields

or equivalently

This means Pis +1is  i2i1 = Pis +1  i2 Pis  i1 Pis  i2 , which is exactly the (s − 1)th-order factorization conjecture. Equation (18.28) also leads to PAis …i2 J Ais …i2 A J Ais …i2 B J~ Ais …i2 * J Ais …i2 = = = = PBis …i2 J Bis …i2 A J Bis …i2 B J~ Bis …i2 * J Bis …i2

(18.30)

which is exactly the steady-state equations of (s − 1)th-order model.

18.3

DNA REPLICATION: A BINARY COPOLYMERIZATION IN TWO DIMENSIONS

Structural and functional studies show that the molecular motor DNAp has two parts. One is the synthetic domain (i.e., polymerase), which can bind the incoming dNTP and catalyze its incorporation into the nascent ssDNA strand (called as primer below for convenience) and at the same time discriminate between the incorporation rates of matched and mismatched nucleotides simply by sensing their different geometry inside the catalytic pocket [5]. Another is the proofreading domain (i.e., exonuclease), which much likely excises the just-incorporated mismatched nucleotide in the primer once the mismatched terminus slips from the polymerase site into the exonuclease site. Thus, DNA replication is regarded approximately as a binary copolymerization process of matches (denoted as A) and mismatches (denoted as B) in two dimensions (polymerase and exonuclease). The first model that explicitly invokes the exonuclease, referred to as Galas-Branscomb model in here, was proposed by Galas and Branscomb [40] and revisited by many other groups [41–44]. Many experimental studies gave consistent results to this model [6,45,46]. In recent years, improved experimental techniques revealed more details of the synthesizing and proofreading processes [8,47], and several detailed kinetic models have been proposed [8,48,49]. However, all these models are based on the original simple Galas-Branscomb model, and many important details such as higher-order neighbor effects at the primer terminus are not considered systematically [49]. In particular, recent experimental works on phi29 DNA polymerase [50,51] revealed more details about the working mechanism of DNAp, highlighting the importance of the forward and backward translocation steps, which are totally absent from the Galas-Branscomb model. Considering this point, as well as many other structural [52–56] and kinetic [6,43,50,51,57] experimental results, we show the full reaction scheme of DNAps that possesses both polymerase site and exonuclease site in Figure 18.3. There are several key features in this comprehensive scheme.

Kinetic Study of the Fidelity of DNA Replication

193

FIGURE 18.3 The reaction scheme for the two-site DNAp. E denotes the enzyme DNA polymerase, and Dn denotes the state of the primer, with n being the length of the primer. Ep·Dn and E′p Dn denote the ‘polymerase-type’ complex when DNAp is in the pre-translocation state or post-translocation state, respectively. Correspondingly, Ex·Dn and E x′ Dn are the two ‘exonuclease-type’ substrates as mentioned in the main text. A free dNTP can bind to DNAp when the complex is at the post-translocation state E′p Dn. When the dNTP is incorporated into the primer, the complex will return to the pre-translocation state Ep Dn+1. The primer terminus may be unzipped from the duplex and slip into the exonuclease site. Model I and Model II are two possible pathways after the nucleotide cleavage in the exonuclease site [12].

First, dsDNA containing a template strand and a primer strand bind to DNAp and form two types of complexes. In the ‘polymerase type’, the template and the primer are paired as duplex and the 3’ terminal of the primer is in the polymerase site. In the ‘exonuclease type’, the template still threads through the polymerase site but the 3’ terminal of the primer is in the exonuclease site. For the ‘polymerase-type’ complexes, there are at least two substrates [50,51]. One is the pre-translocation state of DNAp in which the dNTP binding site is occupied by the primer terminus. Another is the post-translocation state in which the DNAp translocates forward (relative to the template) to expose the site to bind the next dNTP. DNAp can switch between these two states. Correspondingly, one can also assume two substrates of DNAp in the ‘exonuclease-type’ complexes, though there are not sufficient experimental evidences. One is the post-translocation state in which the exonuclease site has been emptied, while the primer terminus does not return to the polymerase site. The other is the pre-translocation state in which the DNAp translocates backward (relative to the template) to force the new primer terminal to occupy the exonuclease site. Second, once the incoming dNTP is incorporated into the primer, the DNAp can either translocate forward to bind a new dNTP in the polymerase site, or it pauses and the primer terminus is peeled off from the duplex and slips into the exonuclease site (the primer terminus can switch backand-forth between the two sites without being excised [51]). The large distance about 30 ∼ 40 Å [52–56] between the two sites implies that the more than one nucleotides of the primer terminus must be unzipped, and thus, the stability of the duplex terminus may put an impact on the slippage probability of the primer terminus. In other words, the more mismatches are incorporated in the primer terminus, the more probable that the terminus slips into the exonuclease site, and the larger probability that mismatches can be proofread. Such higher-order neighbor effects can be significant for the replication fidelity [58] and should be taken account of in the kinetic models, which are totally absent in previous models (for details, see later sections). Third, the exonuclease site can only excise the terminal nucleotide. What happens after the cleavage is not clear yet [59]. Here, we propose two possible pathways, which are denoted as Model I and Model II in Figure 18.3. In Model I, DNAp undergoes a backward translocation and the primer

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Biomotors and their Nanobiotechnology Applications

FIGURE 18.4 The simplified reaction scheme. Xs, Xe: pre-translocation state of DNAp when the primer terminus is in the synthetic(s) site or the exonuclease(s) site, respectively; Xs´, Xe´: post-translocation state of DNAp. It should be noted that when the primer terminus is in the exonuclease site, one does not need to distinguish between ∼ Ae(∼ Be). However, it is still convenient to use ∼ Ae(∼ Be) to denote the immediate state when the terminus switches back to the polymerase site. By setting all the excision rates equal to re, we obtain the models for real DNAp. Under the steady-state condition, the effective rate of dNTP addition can be expressed as f Xs1 = kp[X1], where kp is an effective quasi-first-order rate constant, and [X1] is the concentration of the incoming matched (X1 = A) or mismatched (X1 = B) nucleotide [12].

terminus can either be excised processively, or transfer back to the polymerase site to situate at the pre-translocation state. In Model II, the primer terminus directly transfers back to the polymerase site and situates at the post-translocation state. Higher-order neighbor effects can exist in any step like dNTP binding, the terminus transfer, etc., for both Model I and Model II. Figure 18.3 can be further simplified into Figure 18.4, considering that incorporation of dNTP in the polymerase site is almost irreversible (i.e., the product PPi of the polymerization reaction is often released irreversibly under physiological conditions). The kinetic parameters in this figure are effective parameters, which are combinations of the original rate constants in Figure 18.3. We will only discuss Model I here.

18.3.1

basiC tHeoRy oF steady-state KinetiCs oF tHe exonuClease pRooFReading Model

It has been shown that the terminal mismatch and even the penultimate mismatch at the primer terminus will greatly reduce the incorporation rate of the next nucleotide, compared with the case that a match is at the same position [6,43,57,58,60]. This means that some rate constants in Figure 18.3 are dependent on the few consecutive base pairs at the terminal region, i.e., there do exist higherorder terminal effects in DNA replication by DNAp. Thus, the zero-order terminal model shown in Figure 18.4 is not appropriate, and higher-order models are required. It should be noted first of all that each step in the reaction scheme in Figure 18.3 may have terminal effect but of different order. For instance, the polymerase rate may be of first order, while the transfer rate may be of zero order. This is actually a limiting case of the general first-order scheme shown in Figure 18.5 simply by se se putting k AX 1 = k BX1 and so on. Similarly, schemes with kinetic parameters up to sth order can be included in the general sthorder scheme. 18.3.1.1 First-Order Proofreading Model In this subsection, we will discuss the general first-order proofreading model shown in Figure 18.5 to demonstrate the basic ideas of our approach. Following the same logic of Section 18.2, we use PXsn  X1 to denote the occurrence probability of the terminal sequence X n  X1 in the synthetic (or polymerase) site, PXen  X1 to denote the occurrence probability of Xn…X1 in the exonuclease site, X i = A, B . N Xn  X2 X1 is defined as the number of sequence X n  X 2 X1 in the entire primer chain.

195

Kinetic Study of the Fidelity of DNA Replication

FIGURE 18.5

The minimal scheme of the first-order proofreading model [12].

The overall incorporation rate of sequence X n  X 2 X1 (n ≥ 2) is defined as N. Xn … X2 X1 ≡ J Xn … X2 X1 = J Xs n … X2 X1 + J xen … X2 X1 where J Xs n  X2 X1 ≡ f Xs2 X1 PXsn  X2 , J Xe n  X2 X1 = −rXe2 X1 PXen  X2 X1 The kinetic equations of PXmn  X2 X1 (n ≥ 1, m = s, e) can be written as below P.Xsn … X2 X1 = J Xs n … X2 X1 − J~ Xs n … X2 X1 * − J Xsen … X2 X1

(18.31)

P.Xen … X2 X1 = J Xe n … X2 X1 − J~ Xe n … X2 X1 * − J Xsen … X2 X1

(18.32)

where J~Xs n…X1* = J Xs n…X1 A + J Xs n…X1B , J~ Xe n…X1* = J Xe n…X1 A + J Xe n…X1B , J Xsen…X2 X1 = k Xse2 X1 PXsn…X2 X1 − k Xes2 X1 PXenX2 X1 For example, s s = f AB PAB ( PAAs + PBAs ) − ( fBAs + fBBs ) PABs − k ABse PABs + k ABes PABe s s s + J ABB = J AB − ( J ABA ) − J ABse , e e e e e e e se s es e PAB = −rAB PAB + rBA PABA + rBB PABB + k AB PAB − k AB PAB e e e = J AB − ( J ABA + J ABB ) + J ABse .

(18.33)

(18.34)

What is concerned in this chapter is the steady-state kinetics, i.e., P.Xsn … X2 X1 = 0 and P.Xen … X2 X1 = 0. To rigorously solve these coupled equations, we extend the logic of Section 18.2 and propose the following factorization conjecture of the chain-end sequence distribution. n

m X n  X 2 X1

P

=

∏P

s X i X i −1

i= 3

  

n

∏P

s X i −1

i =3

−1

  PXm2 X1 , n ≥ 3, m = s, e. 

which means that the symbolic sequence of the growing primer can be described as a first-order Markovian chain, which is generated in the polymerase pathway, and the existence of exonuclease pathway does not change this Markovian characteristics. For example, PXs3 X2 X1 = PXs3 X2 PXs2 X1 PXs2 , PXe3 X2 X1 = PXs3 X2 PXe2 X1 PXs2 . The validity of the factorization conjecture has been tested by Monte-Carlo simulation as shown in Figure 18.6.

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FIGURE 18.6 Simulation verification of the factorization conjecture of the first-order model, with illustras s s s e e e e se se se tive rate parameters f AA  = 8, f AB  = 6, f BA  = 4, f BB  = 2, rAA  = 1, rAB  = 1, rBA  = 2, rBB  = 1, k AA  = 1, k AB  = 6, k BA  = 1, se es es es es k BB  = 6, k AA  = 1, k AB  = 3, k BA  = 1, k BB  = 4. Averaged over 10,000 samples [12].

By this factorization conjecture, the original unclosed equations can be reduced to the following closed equations for the eight basic variables PXm2 X1 (m = s, e). e e − J AB = J Bse , J BA

s s J BA − J AB = J Ase ,

s se s s se s J AA − J AA PAA J AB − J AB PAB , , = = s se s s se s J BA − J BA PBA J BB − J BB PBB e se se + J AA Ps J e + J AB Ps J AA , AB , = AA = AB e se s e se s J BA + J BA PBA J BB + J BB PBB

J Ase + J Bse = 0

∑

(P

s XY

(18.35)

e + PXY )=1

X ,Y = A, B

18.3.1.2 Second-Order Proofreading Model Second-order terminal effects have been observed for some DNAps where the penultimate mismatch at the terminus can affect the next nucleotide incorporation [58,60]. In this subsubsection, we extend the method of the preceding subsubsection and discuss the second-order proofreading model as shown in Figure 18.7. Similar to the first-order model, we have N. Xn … X2 X1 ≡ J Xn … X2 X1 = J Xs n … X2 X1 + J Xe n … X2 X1 (n ≥ 3) where J Xs n  X3 X2 X1 = f Xs3 X2 X1 PXsn  X3 X2 , J Xe n  X3 X2 X1 = −rXe3 X2 X1 PXen  X3 X2 X1 The kinetic equations for PXmn  X3 X2 X1 (n ≥ 1, m = s, e) can be written as below

P.Xsn … X3 X2 X1 = J Xs n … X3 X2 X1 − J~Xs n … X3 X2 X1 * − J Xsen … X3 X2 X1 ,

(18.36)

P.Xen … X3 X2 X1 = J Xe n … X3 X2 X1 − J~Xe n … X3 X2 X1 * + J Xsen … X3 X2 X1

(18.37)

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Kinetic Study of the Fidelity of DNA Replication

FIGURE 18.7

The second-order proofreading reaction scheme [12].

where J Xsen  X3 X2 X1 = k Xse3 X2 X1 PXsn  X3 X2 X1 − k Xes3 X2 X1 PXen  X3 X2 X1 Under the steady-state conditions P.Xsn … X3 X2 X1 = P.Xen … X3 X2 X1 = 0, we proposed the following factorization conjecture: n

m X n  X 3 X 2 X1

P

=

∏P

s X i X i −1 X i − 2

i= 4

  

n

∏P

s X i −1 X i − 2

i=4

−1

  PXm3 X2 X1 n ≥ 4, m = s, e 

which can also be tested by Monte-Carlo simulations (results not shown here). Therefore, we obtain the following closed equations for the second-order proofreading model s s se − JXX* = J XX , J XX

s s se J XXX − J XXX = J XX ,

s se J AXY − J AXY Ps = AXY , J XXX = J XXX , s se s J BXY − J AXY PBXY e se + J AXY Ps J AXY , J AB = J BA , = AXY s e se J BXY + J BXY PBXY s e + PXYZ ∑ ( PXYZ ) = 1, X ,Y , Z = A, B,

where X is A(B) and X is different from X. Some experiments [43,61] show that up to four base pairs at the primer terminus may have apparent effects on the incorporation of the next nucleotide. For such cases, one should generalize the above method to include these higher-order terminal effects. The generalization to sth-order model is straightforward (details not given here).

18.3.2

tHe Fidelity oF dna RepliCation

In this subsection, we discuss the fidelity problem of molecular motor DNAp in details. In principle, one can define the fidelity naturally as the ratio of matches over mismatches incorporated into the primer during the replication processes under steady-state conditions. However, it is difficult to directly measure this fidelity in experiments, and some indirect methods were developed. One of

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the commonly used methods is the forward mutation assay [46,62–64], which scores the replication errors indirectly by counting the phenotype change rate of the bacterial hosts transfected by reporter gene DNA, or sometimes by directly sequencing the mismatched base pairs in the replicated DNA. Other frequently used methods are steady-state [65–67] or pre-steady-state [6,68–70] kinetic assays, which studied the kinetics of DNA replication directly (i.e., the reaction pathway and the corresponding rate constants, in particular, rates of match or mismatch incorporation), and calculate the replication fidelity indirectly based on the theoretical models. The basic principles of these two approaches differ, but the calculated fidelity is often of similar magnitudes. For example, the average fidelity of Sulfolobus solfataricus P2 DNA polymerase IV (Dpo4) is about 1.3 × 102 to 3.3 × 103 using the steady-state kinetic method [71], which agrees with 1.5 × 102 given by the forward mutation assay [63]. In general, for most proofreading-proficient DNAps, the fidelity in vitro is about 106 ∼ 107 with a contribution by exonuclease proofreading of 101 ∼ 102 [5,72]. In this subsection, we will only discuss the kinetics-based fidelity, since it can be rigorously defined and calculated within the framework of our basic theory. It is worth pointing out that previous works (e.g., Refs. [73,74]) relating kinetic parameters to fidelity were done approximately by using simple methods like Michaelis-Menten kinetics, which are in fact invalid for exonuclease proofreading, as pointing out in preceding subsections. Additionally, higher-order terminal effects will be a focus of this subsection. In our theoretical framework, the replication fidelity is expressed as φ = NA /NB. Here, NA is the total number of incorporated matches in the primer, and NB is the total number of mismatches. Once the closed equations in the preceding sections are solved, the total flux JA, JB can be calculated. Since N A = J A , N B = J B , and d ( N A N B ) dt = 0 (in steady state), we can calculate the replication fidelity exactly by φ = NA /NB = JA /JB. Furthermore, it will be very useful for experimentalists and further theoretical studies to obtain analytical expressions of φ in terms of the kinetic parameters. However, it is often impossible to solve the algebra equations like Eq. (18.36) analytically. To circumvent this problem, we introduce the below another method referred to as infinite-state Markov chain [37] to calculate φ. This method has already been used for higherorder copolymerization in Section 18.2. Below, we show that the same logic can be extended to the exonuclease proofreading reaction scheme, and this method does give exactly the same numerical results of φ as that given by our steady-state kinetic equations (data not shown). From this aspect, both methods serve as a mutual verification of each other. Besides, the mathematical expression given by the infinite-state Markov chain method can be further simplified into intuitive forms under some biologically relevant conditions (see below) to help us to appreciate which parameters play the key role in determining the fidelity and how they do it, in particular in terms of high-order terminal effects. 18.3.2.1 The Infinite-State Markov Chain Method for Exonuclease Proofreading We begin with the first-order reaction scheme. The branching model for this case is illustrated in Figure 18.8. This growing chain is completely characterized by four groups of transition probabilities, which can be expressed as PX X2 X1 ≡ f Xs1X PXse2 X1 ≡ k Xse2 X1

(f (f

s X1 X

s X1 X

) ),

+ f Xs1Xˆ + k Xse2 X1 , + f Xs1Xˆ + k Xse2 X1

PXes2 X1 ≡ k Xes2 X1 ( rXe2 X1 + k Xes2 X1 ) , Pu X2 X1 ≡ 1 − PXes2 X1 = rXe2 X1 ( rXe2 X1 + k Xes2 X1 )

(18.38)

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Kinetic Study of the Fidelity of DNA Replication

FIGURE 18.8 Branching model for the first-order polymerization and excision [12].

We also employ the idea of ‘cycle completion’ [37]. In the case here, it is that, when a nucleotide is incorporated, it has a chance to be excised, only those not being excised account for the final composition. Thus, the fidelity for the first-order terminal model can be defined as

φ=

QAA + QBA QAB + QBB

(18.39)

where QX2 X1 is the probability of eventually incorporating X1 to the terminal X2, satisfying QAA + QAB + QBA + QBB = 1. QX2 X1 can be explicitly expressed as QX2 X1 ≡ PˆX2 X1 PnuX2 X1 , where PˆX2 X1 is the probability that incorporating X1 to the terminal X 2 , PnuX2 X1 is the probability of the terminal X2 X1 never being excised. For the first-order model, the absolute values of PˆX2 X1 are not known a prior, but the following equalities obviously hold: s PA AB PA BB Pˆ fs PˆAA PA AA PA BA f AA = S , BA = = = = = BA s f BB PˆAB PB AA PB BA f AB PˆBB PB AB PB BB

(18.40)

Considering the fact that the number of AB should equal to the number of BA in the copolymer chain, we have the following intrinsic constraint:

(

)

(

QAB = PˆAB PnuAB = QBA = PˆBA PnuBA

)

(18.41)

To calculate PnuX2X1, we define PeuX2X1 ≡ 1 − PnuX2X1 as the probability of the terminal X2 X1 ever being excised. PeuX2X1 satisfy the following iterative equations (details can be found in Ref. [12]):

PeuX2 X1 =

Pˆu X2 X1 se PX2 X1 PXes2 X1

  1 1   −  1 − PˆA X2 X1 PeuX1 A + PˆB X2 X1 PeuX1B PXes2 X1 TX2 X1 

(

)

(18.42)

Here, TX2 X1 = 1 (1 − PXse2 X1 PXes2 X1 ) , Pˆu X2 X1 = Pu X2 X1 pXse2 X1 TX2 X1 = rˆXs2 X1 ( f Xs1 A + f Xs1B + rˆXs2 X1 ) , PˆX X2 X1 = PX X2 X1 TX2 X1 = f Xs1X f Xs1X + f Xs1Xˆ + rˆXs2 X1 , and rˆXs2 X1 = k Xse2 X1 rXe2 X1 ( rXe2 X1 + k Xse2 X1 ) .

(

)

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Biomotors and their Nanobiotechnology Applications

Once PeuX2 X1 are solved, PˆX2 X1 can then be calculated by combining Eqs. (18.40) and (18.41), and the fidelity φ can be obtained by Eq. (18.39). φ calculated in this approach is found numerically identical to that given by the steady-state kinetic equations, which can be regarded as a verification of our kinetic approach. The same logic can be extended to any higher-order models. 18.3.2.2 Approximation of φ under Bio-Relevant Conditions The kinetic parameters of DNAp have several common features. For example, match incorporation at polymerase site is always very fast, and also much faster than mismatch incorporation. The existence of large difference in order of magnitude of the kinetic parameters enables us to make some reasonable approximations (called bio-relevant conditions) to simplify the above procedure to obtain the explicit mathematical expression of φ in terms of several key parameters. It should be noticed first that the subscript of A and B for the excision step is meaningless; thus, one can use re to denote all the rXes +1 Xs  X1 . In Table 18.1, we list experimental values of some kinetic parameters for some DNAps. Inspired by these data, we propose the following bio-relevant conditions of kinetic parameters for the sth-order model. s s a. f AAA… >> f AAA …B , which leads to PA| X AAA... >> PB| X AAA... h +1

h

h

h

s b. f AAA… >> k XseAAA… (> rˆXs AAA... ) , which leads to PA| X AAA... >> PXseAAA… (> Pˆu| X AAA... ) _ _ h +1

h

h

h

h

h

TABLE 18.1 Experimental Value of Kinetic Parameters (s−1) of Some Real DNAps T7 [7,57]

Pol γ [58,60]

f

s AA

250a

3,900–5,700c

f

s AB

0.002a

0.023–1.6c

se f AA

0.2b

>0.05d

0.015f

f

se AB

2.3

b

>0.4

0.038

5

–

f

es AA

714b

>39d

–

20

10.48i

es f AB

714b

–

–

–

–

e

896

>39

100

500h

se k ABA

–

>3

–

–

–

es ABA

–

–

–

–

–

s f ABA

0.012a

0.1c

–

–

10−3–10−4g

s f BAA

–

2.7c

–

–

–

Rate Parameter First order

r Second order

k

b

d

d

d

Pol III [75,76] 370e (0.16–2.1) × 10

−3e

f

280

f

T4 [77]

phi29 DNAP [51,78,79]

600

680g

∗

10−4–10−6g

1

11.54i

All forward polymerization kinetic parameters are scaled to the standard dNTP concentration 100 μM. – means the data cannot be found. ∗ means the data is too small to measure. a From Table II of Ref. [7]. b From Ref. [57]. c From Ref. [60]. dNTP concentration is set as 100 μM for holoenzyme. d Estimated from the combined kinetic parameters from Ref. [58]. e Values for the holoenzyme, from Ref. [76]. f Estimated by pre-steady-state measurements of purified ε subunit, from Ref. [75]. g From Table I and Table II of Ref. [78] in Mg2+-activated polymerization. h From Ref. [79]. i From Ref. [51].

201

Kinetic Study of the Fidelity of DNA Replication

s This can be easily achieved by tuning the concentration [A] (notice that f AAA… is proporh+1

tional to [A]). The matched terminus will continue to incorporate the next match, instead of being transferred to the exonuclease site. s s ˆ ˆ c. rˆAAA...B > f AAA...B AAA… AAA... for 0 < i ≤ m (which leads to Pu|AAA...B AAA… (≡ Ri ) > PA|AAA...B AAA… (≡ Fi ), _ _ i −1

h − i +1

∼

h − i +1

i

h−1

∼

i −1

h −i +1

i −1

s s and rˆAAA...B AAA… rˆAB f_ k se_ _ B , which leads to PA| AAA… P _ _ PAAA…