Bacterial and Archaeal Motility 1071630598, 9781071630594

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Methods in Molecular Biology 2646

Tohru Minamino Makoto Miyata Keiichi Namba Editors

Bacterial and Archaeal Motility

METHODS

IN

MOLECULAR BIOLOGY

Series Editor John M. Walker School of Life and Medical Sciences University of Hertfordshire Hatfield, Hertfordshire, UK

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

For over 35 years, biological scientists have come to rely on the research protocols and methodologies in the critically acclaimed Methods in Molecular Biology series. The series was the first to introduce the step-by-step protocols approach that has become the standard in all biomedical protocol publishing. Each protocol is provided in readily-reproducible step-bystep fashion, opening with an introductory overview, a list of the materials and reagents needed to complete the experiment, and followed by a detailed procedure that is supported with a helpful notes section offering tips and tricks of the trade as well as troubleshooting advice. These hallmark features were introduced by series editor Dr. John Walker and constitute the key ingredient in each and every volume of the Methods in Molecular Biology series. Tested and trusted, comprehensive and reliable, all protocols from the series are indexed in PubMed.

Bacterial and Archaeal Motility Edited by

Tohru Minamino Graduate School of Frontier Biosciences, Osaka University, Suita, Osaka, Japan

Makoto Miyata Graduate School of Science, Osaka Metropolitan University, Osaka, Japan

Keiichi Namba Graduate School of Frontier Biosciences, Osaka University, Suita, Osaka, Japan

Editors Tohru Minamino Graduate School of Frontier Biosciences Osaka University Suita, Osaka, Japan

Makoto Miyata Graduate School of Science Osaka Metropolitan University Osaka, Japan

Keiichi Namba Graduate School of Frontier Biosciences Osaka University Suita, Osaka, Japan

ISSN 1064-3745 ISSN 1940-6029 (electronic) Methods in Molecular Biology ISBN 978-1-0716-3059-4 ISBN 978-1-0716-3060-0 (eBook) https://doi.org/10.1007/978-1-0716-3060-0 © The Editor(s) (if applicable) and The Author(s), under exclusive license to Springer Science+Business Media, LLC, part of Springer Nature 2023 This work is subject to copyright. All rights are solely and exclusively licensed by the Publisher, whether the whole or part of the material is concerned, specifically the rights of translation, reprinting, reuse of illustrations, recitation, broadcasting, reproduction on microfilms or in any other physical way, and transmission or information storage and retrieval, electronic adaptation, computer software, or by similar or dissimilar methodology now known or hereafter developed. The use of general descriptive names, registered names, trademarks, service marks, etc. in this publication does not imply, even in the absence of a specific statement, that such names are exempt from the relevant protective laws and regulations and therefore free for general use. The publisher, the authors, and the editors are safe to assume that the advice and information in this book are believed to be true and accurate at the date of publication. Neither the publisher nor the authors or the editors give a warranty, expressed or implied, with respect to the material contained herein or for any errors or omissions that may have been made. The publisher remains neutral with regard to jurisdictional claims in published maps and institutional affiliations. This Humana imprint is published by the registered company Springer Science+Business Media, LLC, part of Springer Nature. The registered company address is: 1 New York Plaza, New York, NY 10004, U.S.A.

Preface Many bacteria and archaea migrate toward more favorable environments for their survival by using their own motility machinery. Motility is also required for effective infection of pathogenic bacteria into their host cells. Thus, bacterial and archaeal motility is an extremely intriguing topic. Many motile bacteria utilize flagella for their swimming motility in liquid environments and their swarming motility on solid surfaces. The bacterial flagellum is a motility organelle driven by a rotary motor powered by the transmembrane electrochemical gradient of ions such as protons (H+) and sodium ions (Na+). Motile archaea also have a flagellum-like structure called the archaellum, which looks very similar to the bacterial flagellum both in appearance and function. In contrast to bacterial flagella, however, ATP is the primary energy source to rotate the archaellum. Many bacteria also utilize the ATP-driven type IV pilus for twitching motility over surfaces. Interestingly, Mollicutes species have neither flagella nor type IV pili and use their own ATP-driven motility machinery for gliding motility on surfaces and swimming motility in liquids. Thus, bacterial and archaeal motility can be defined as the capability of individual cells to convert electrochemical or chemical energy to mechanical work required for their locomotion under various environmental conditions. In this volume, we have brought together a set of cutting-edge research protocols to study the structure and dynamics of bacterial and archaeal motility systems using bacterial genetics, molecular biology, biochemistry, biophysics, structural biology, cell biology, microscopy imaging, and molecular dynamics simulation. Our aim is to provide useful tools and pathways for the investigation of the supramolecular motility machines derived from various bacterial and archaeal species through techniques that can be applied. Since the principal goal of the book is to provide researchers with a comprehensive account of the practical steps of each protocol, the Methods sections contain detailed step-by-step descriptions of every protocol. The Notes sections complement the Methods to get the hang of each experiment based on the authors’ experiences and to figure out the best way to solve any problem and difficulty that might arise during the experiment. The bacterial flagellum is a supramolecular assembly composed of 30 different proteins and consists of at least 3 structural parts: a basal body that acts as a bi-directional rotary motor, a filament that works as a molecular screw to produce thrust to propel the cell body, and a hook that connects the basal body and filament and serves as a universal joint to transmit torque produced by the motor to the molecular screw. Bacteria employ the flagellar type III secretion systems (fT3SS) to export flagellar proteins to construct the flagella on the cell surface. The fT3SS is composed of a transmembrane export gate complex and a cytoplasmic ATPase ring complex. The fT3SS utilizes free energy derived from ATP hydrolysis by the cytoplasmic ATPase complex and proton motive force across the cytoplasmic membrane to unfold and transport flagellar structural subunits from the cytoplasm to the distal end of the growing flagellar structure where each subunit is folded back and incorporated into the structure. Chapters in Part I describe how to purify the core complex of the export gate complex (Chapter 1), how to quantitatively measure flagellar protein export in vitro (Chapter 2), how to visualize the assembly and ejection processes of the flagella (Chapter 4), and how to carry out high-resolution cryoEM structural analysis of the flagellar filament (Chapter 5). Molecular dynamic simulation of dynamic open-close domain

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motion of a transmembrane export gate protein responsible for efficient and robust energy coupling mechanism of the fT3SS (Chapter 3) is also included in Part I. The bacterial flagellar motor consists of a rotor and a dozen stator units. The energy for motor rotation is supplied by ion influx along the electrochemical potential difference of specific ions, such as H+ and Na+, across the cytoplasmic membrane. The stator unit is composed of two transmembrane proteins, commonly referred to as MotA and MotB in the H+-driven motor, PomA and PomB in the Na+-driven motor of marine Vibrio, and MotP and MotS in the Na+-driven motor of extremely alkalophilic Bacillus. The stator unit serves also as a transmembrane ion channel complex to couple the ion flow through the channel with torque generation via electrostatic interactions of either MotA, PomA, or MotP with a rotor protein, FliG. The flagellar motor can spin in both counter-clockwise (CCW, viewed from outside) and clockwise (CW) directions without changing the direction of the ion flow. Several chemotaxis signaling molecules bind to the rotor and induce its highly cooperative conformational changes. As a result, the flagellar motor switches its rotational direction from CCW to CW. When the signaling molecules dissociate from the rotor, the motor spins again in the CCW direction. Chapters in Part II will describe how to analyze the structural dynamics of FliG (Chapter 6), how to investigate stator-rotor interactions in vivo (Chapter 7), how to measure the ion channel activity of the stator unit (Chapters 8 and 9), how to visualize structural dynamics of the stator unit in vitro (Chapter 10), and how to measure flagellar motor dynamics at near zero load (Chapter 11). Flagella-driven magnetotaxis motility (Chapter 12), flagella-driven surface motility of Salmonella (Chapter 13), and flagella-driven motility of Leptospira (Chapters 14 and 15) are also included in Part II. Motile archaea rotate long helical filaments called the archaella to swim in liquid environments. The archaella look like the bacterial flagella, but the archaellar structure has nothing in common with the bacterial flagella. The rotating appendage is formed by a long, helical filament, which is stably attached to a membrane-embedded rotary motor powered by ATP hydrolysis. The N-terminus of archaellar filament proteins contains a cleavable signal peptide that is removed during secretion, and each filament subunit is incorporated at the proximal end of a nascent archaellar structure in a way similar to the type IV pilus. Furthermore, the cytoplasmic ATPase ring complex, which is responsible for both assembly and rotation, is structurally and functionally similar to the type IV pilus ATPases, suggesting that the archaellum may have been evolved from the type IV pilus. Chapters in Part III describe how to isolate the archaellar filaments (Chapter 16) and how to measure ATP-driven archaellar motor rotation (Chapter 17). Bacteria utilize type IV pili to move over surfaces, and this crawling bacterial motility is called twitching motility. Each type IV pilus is composed of a basal body located within the cell envelope and a pilus filament extended from the cell body. The pilus filament is a highly dynamic structure that undergoes cycles of extension and retraction. Two distinct ATP-driven motors located at the base of the filament are directly involved in such dynamic cycles of the pilus filament motion. One ATP-driven motor facilitates the extension of the pilus filament, allowing the filament to attach to the surface of the host cells and substrates, and then the other ATP-driven motor promotes pilus retraction, resulting in the forward movement of the cell body. Chapters in Part IV describe how to perform structural (Chapter 18) and functional analyses (Chapters 19 and 20) of type IV pili of twitchingcapable bacteria. Some bacterial species have their own specialized motility machinery to drive gliding motility over surfaces. This type of bacterial motility does not involve a conventional bacterial motility machinery such as flagella and type IV pili. Bacteroidota adhere to solid

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surfaces through their own adhesions and move back and forth by the movement of the adhesins on helical filamentous structures located in the periplasm. Adhesive movements are dependent on proton motive force across the cytoplasmic membrane. The mechanical force required for adhesive migration should occur around the cytoplasmic membrane and be transmitted to the outer membrane surface via the peptidoglycan layer. Chapters in Part V describe how to perform structural (Chapter 21) and functional analyses (Chapters 22 and 23) of the gliding motility machinery of Bacteroidota. A chapter describing how to visualize the peptidoglycan layer structures (Chapter 24) is also included in Part V. Mycoplasma species utilize adhesins to catch, pull, and release sialylated oligosaccharides on the host cell surfaces to drive gliding motility over the cell surfaces. The gliding motility of Mycoplasma is powered by ATP hydrolysis. Thus, the gliding motility machinery of Mycoplasma is structurally and functionally different from that of Bacteroidota. Interestingly, the structure and locomotion mechanism of the ATP-driven gliding motility machinery of Mycoplasma pneumoniae and Mycoplasma mobile, both of which belong to the class Mollicutes, seem to be different from each other. Furthermore, Spiroplasma, which also belong to the class Mollicutes, utilize a unique motility machinery to swim in liquid environments. Chapters in Part VI describe how to carry out structural (Chapter 25) and functional analyses of the motility machinery of Mollicutes (Chapters 29 and 30) and how to directly measure the velocity and force generated by the motility machinery of Mollicutes using high-resolution optical microscopic techniques (Chapters 26, 27, 28, and 31). Chloroflexus has its own gliding motility machinery, but the machinery is not yet identified. A chapter describing how to characterize the gliding motility of Chloroflexus (Chapter 32) is also included in Part VI. All the contributors are leading researchers in the field of bacterial and archaeal motility, and we would like to acknowledge them for providing their comprehensive protocols and techniques for this volume. We would like to thank Dr. John Walker, Editor-in-Chief of the Methods in Molecular Biology series, for giving us a great opportunity to edit this volume and his continuous support and encouragement. We hope you all enjoy these chapters and benefit from them in this volume of Methods in Molecular Biology. Osaka, Japan Osaka, Japan Osaka, Japan

Tohru Minamino Makoto Miyata Keiichi Namba

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

PART I

BACTERIAL FLAGELLAR PROTEIN EXPORT AND ASSEMBLY

1 Purification of the Transmembrane Polypeptide Channel Complex of the Salmonella Flagellar Type III Secretion System . . . . . . . . . . . . . . . . . . . . . . . Miki Kinoshita, Keiichi Namba, and Tohru Minamino 2 In Vitro Flagellar Type III Protein Transport Assay Using Inverted Membrane Vesicles . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Katsumi Imada and Hiroyuki Terashima 3 Molecular Simulation to Investigate Open–Close Motion of a Flagellar Export Apparatus Protein FlhAC . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Akio Kitao 4 Live-Cell Imaging of the Assembly and Ejection Processes of the Bacterial Flagella by Fluorescence Microscopy . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Xiang-Yu Zhuang, Chao-Kai Tseng, and Chien-Jung Lo 5 Purification and CryoEM Image Analysis of the Bacterial Flagellar Filament . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Tomoko Yamaguchi, Tomoko Miyata, Fumiaki Makino, and Keiichi Namba

PART II

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3

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27

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43

FLAGELLA-DRIVEN MOTILITY OF BACTERIA

6 Site-Specific Isotope Labeling of FliG for Studying Structural Dynamics Using Nuclear Magnetic Resonance Spectroscopy. . . . . . . . . . . . . . . . . . . . . . . . . . . 57 Tatsuro Nishikino and Yohei Miyanoiri 7 Site-Directed Cross-Linking Between Bacterial Flagellar Motor Proteins In Vivo . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 71 Hiroyuki Terashima, Michio Homma, and Seiji Kojima 8 Measurements of the Ion Channel Activity of the Transmembrane Stator Complex in the Bacterial Flagellar Motor . . . . . . . . . . . . . . . . . . . . . . . . . . . . 83 Yusuke V. Morimoto and Tohru Minamino 9 Purification of the Na+-Driven PomAB Stator Complex and Its Analysis Using ATR-FTIR Spectroscopy. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 95 Seiji Kojima, Michio Homma, and Hideki Kandori 10 Purification of Na+-Driven MotPS Stator Complexes and Single-Molecule Imaging by High-Speed Atomic Force Microscopy . . . . . . . . . . . . . . . . . . . . . . . . . 109 Naoya Terahara and Noriyuki Kodera 11 High-Resolution Rotation Assay of the Bacterial Flagellar Motor Near Zero Loads Using a Mutant Having a Rod-Like Straight Hook . . . . . . . . . . . . . . 125 Shuichi Nakamura and Tohru Minamino

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Live-Cell Fluorescence Imaging of Magnetosome Organelle for Magnetotaxis Motility . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Yukako Eguchi and Azuma Taoka 13 Swarming Motility Assays in Salmonella . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Jonathan D. Partridge and Rasika M. Harshey 14 Analysis of Adhesion and Surface Motility of a Spirochete Bacterium . . . . . . . . . . Shuichi Nakamura, Jun Xu, and Nobuo Koizumi 15 Force Measurement of Bacterial Swimming Using Optical Tweezers . . . . . . . . . . Keigo Abe, Kyosuke Takabe, and Shuichi Nakamura

PART III 16

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133 147 159 169

ARCHAELLA-DRIVEN MOTILITY OF ARCHAEA

Archaella Isolation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 183 ˜ o N. de Sousa Machado, Shamphavi Sivabalasarma, Joa Sonja-Verena Albers, and Ken F. Jarrell Direct Observation of Archaellar Motor Rotation by Single-Molecular Imaging Techniques. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 197 Yoshiaki Kinosita

PART IV

TYPE IV-DRIVEN TWITCHING MOTILITY OF BACTERIA

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In Situ Structure Determination of Bacterial Surface Nanomachines Using Cryo-Electron Tomography . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 211 Longsheng Lai, Yee-Wai Cheung, Matthew Martinez, Kathryn Kixmoeller, Leon Palao III, Stefan Steimle, Meng-Chiao Ho, Ben E. Black, Erh-Min Lai, and Yi-Wei Chang 19 Twitching Motility Assays of Lysobacter enzymogenes OH11 Under a Light Microscope . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 249 Bingxin Wang, Xiaolong Shao, and Guoliang Qian 20 Live Cell Imaging of the Twitching Motility of Cyanobacteria by High-Resolution Microscopy . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 255 Daisuke Nakane

PART V ADHESION-BASED GLIDING MOTILITY OF BACTERIA Isolation and Visualization of Gliding Motility Machinery in Bacteroidota . . . . . Satoshi Shibata and Daisuke Nakane 22 Live Cell Imaging of Gliding Motility of Flavobacterium johnsoniae Under High-Resolution Microscopy . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Daisuke Nakane and Satoshi Shibata 23 Social Motility Assays of Flavobacterium johnsoniae . . . . . . . . . . . . . . . . . . . . . . . . . Chikara Sato and Keiko Sato 24 Visualization of Peptidoglycan Structures of Escherichia coli by Quick-Freeze Deep-Etch Electron Microscopy . . . . . . . . . . . . . . . . . . . . . . . . . . Yuhei O. Tahara and Makoto Miyata 21

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PART VI 25

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28 29 30 31 32

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UNIQUE MOTILITY SYSTEM IN BACTERIA

Purification and Structural Analysis of the Gliding Motility Machinery in Mycoplasma mobile . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Takuma Toyonaga and Makoto Miyata Motility Assays of Mycoplasma mobile Under Light Microscopy . . . . . . . . . . . . . . . Taishi Kasai and Makoto Miyata Detection of Steps and Rotation in the Gliding Motility of Mycoplasma mobile. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Yoshiaki Kinosita, Mitsuhiro Sugawa, Makoto Miyata, and Takayuki Nishizaka Direct Measurement of Kinetic Force Generated by Mycoplasma . . . . . . . . . . . . . Masaki Mizutani and Makoto Miyata Genetic Manipulation of Mycoplasma pneumoniae . . . . . . . . . . . . . . . . . . . . . . . . . . Tsuyoshi Kenri Purification and ATPase Activity Measurement of Spiroplasma MreB. . . . . . . . . . Daichi Takahashi, Ikuko Fujiwara, and Makoto Miyata Swimming Motility Assays of Spiroplasma. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Daisuke Nakane Motility Assays of Chloroflexus . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Shin Haruta, Hinata Kakuhama, Shun-ichi Fukushima, and Sho Morohoshi

Index . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

311 321

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337 347 359 373 383

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Contributors KEIGO ABE • Department of Applied Physics, Graduate School of Engineering, Tohoku University, Sendai, Miyagi, Japan SONJA-VERENA ALBERS • Molecular Biology of Archaea, Faculty of Biology, Institute of Biology II, University of Freiburg, Freiburg, Germany; Spemann Graduate School of Biology and Medicine, University of Freiburg, Freiburg, Germany BEN E. BLACK • Department of Biochemistry and Biophysics, Perelman School of Medicine, University of Pennsylvania, Philadelphia, PA, USA YI-WEI CHANG • Department of Biochemistry and Biophysics, Perelman School of Medicine, University of Pennsylvania, Philadelphia, PA, USA YEE-WAI CHEUNG • Department of Biochemistry and Biophysics, Perelman School of Medicine, University of Pennsylvania, Philadelphia, PA, USA; Institute of Plant and Microbial Biology, Academia Sinica, Taipei, Taiwan ˜ JOAO N. DE SOUSA MACHADO • Molecular Biology of Archaea, Faculty of Biology, Institute of Biology II, University of Freiburg, Freiburg, Germany; Spemann Graduate School of Biology and Medicine, University of Freiburg, Freiburg, Germany YUKAKO EGUCHI • Institute for Promotion of Diversity and Inclusion, Kanazawa University, Kanazawa, Ishikawa, Japan IKUKO FUJIWARA • Graduate School of Science, Osaka City University, Osaka, Japan; The OCU Advanced Research Institute for Natural Science and Technology (OCARINA), Osaka City University, Osaka, Japan; Department of Bioengineering, Nagaoka University of Technology, Nagaoka, Niigata, Japan; Department of Materials Science and Bioengineering, Nagaoka University of Technology, Nagaoka, Niigata, Japan SHUN-ICHI FUKUSHIMA • SANKEN (Institute of Scientific and Industrial Research), Osaka University, Ibaraki, Osaka, Japan RASIKA M. HARSHEY • Department of Molecular Biosciences, The University of Texas at Austin, Austin, TX, USA SHIN HARUTA • Department of Biological Sciences, Tokyo Metropolitan University, Hachioji, Tokyo, Japan MENG-CHIAO HO • Institute of Biological Chemistry, Academia Sinica, Taipei, Taiwan; Institute of Biochemical Sciences, National Taiwan University, Taipei, Taiwan MICHIO HOMMA • Division of Biological Science, Graduate School of Science, Nagoya University, Nagoya, Japan KATSUMI IMADA • Department of Macromolecular Science, Graduate School of Science, Osaka University, Toyonaka, Osaka, Japan KEN F. JARRELL • Department of Biomedical and Molecular Sciences, Queen’s University, Kingston, ON, Canada HINATA KAKUHAMA • Department of Biological Sciences, Tokyo Metropolitan University, Hachioji, Tokyo, Japan HIDEKI KANDORI • Department of Life Science and Applied Chemistry, Nagoya Institute of Technology, Nagoya, Japan

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Contributors

TAISHI KASAI • College of Science, Department of Life Science, Rikkyo University, Tokyo, Japan TSUYOSHI KENRI • Department of Bacteriology II, National Institute of Infectious Diseases, Musashimurayama, Tokyo, Japan MIKI KINOSHITA • Graduate School of Frontier Biosciences, Osaka University, Suita, Osaka, Japan YOSHIAKI KINOSITA • CPR, RIKEN, Wako, Saitama, Japan AKIO KITAO • School of Life Science and Technology, Tokyo Institute of Technology, Meguro, Tokyo, Japan KATHRYN KIXMOELLER • Department of Biochemistry and Biophysics, Perelman School of Medicine, University of Pennsylvania, Philadelphia, PA, USA NORIYUKI KODERA • Nano Life Science Institute, Kanazawa University, Kanazawa, Japan NOBUO KOIZUMI • Department of Bacteriology I, National Institute of Infectious Diseases, Tokyo, Japan SEIJI KOJIMA • Division of Biological Science, Graduate School of Science, Nagoya University, Nagoya, Japan ERH-MIN LAI • Institute of Plant and Microbial Biology, Academia Sinica, Taipei, Taiwan LONGSHENG LAI • Department of Biochemistry and Biophysics, Perelman School of Medicine, University of Pennsylvania, Philadelphia, PA, USA CHIEN-JUNG LO • Department of Physics, National Central University, Taoyuan, Taiwan; Center for Complex Systems, National Central University, Taoyuan, Taiwan FUMIAKI MAKINO • Graduate School of Frontier Biosciences, Osaka University, Suita, Osaka, Japan; JEOL Ltd., Akishima, Tokyo, Japan MATTHEW MARTINEZ • Department of Biochemistry and Biophysics, Perelman School of Medicine, University of Pennsylvania, Philadelphia, PA, USA TOHRU MINAMINO • Graduate School of Frontier Biosciences, Osaka University, Suita, Osaka, Japan YOHEI MIYANOIRI • Institute for Protein Research, Osaka University, Suita, Osaka, Japan MAKOTO MIYATA • Graduate School of Science, Osaka City University, Osaka, Japan; Graduate School of Science, Osaka Metropolitan University, Osaka, Japan; The OCU Advanced Research Institute for Natural Science and Technology (OCARINA), Osaka City University, Osaka, Japan; The OMU Advanced Research Center for Natural Science and Technology, Osaka Metropolitan University, Osaka, Japan TOMOKO MIYATA • Graduate School of Frontier Biosciences, Osaka University, Suita, Osaka, Japan; JEOL YOKOGUSHI Research Alliance Laboratories, Osaka University, Suita, Osaka, Japan MASAKI MIZUTANI • Bioproduction Research Institute, National Institute of Advanced Industrial Science and Technology (AIST), Tsukuba, Ibaraki, Japan YUSUKE V. MORIMOTO • Department of Physics and Information Technology, Faculty of Computer Science and Systems Engineering, Kyushu Institute of Technology, Iizuka, Fukuoka, Japan; Japan Science and Technology Agency, PRESTO, Kawaguchi, Saitama, Japan SHO MOROHOSHI • Department of Biological Sciences, Tokyo Metropolitan University, Hachioji, Tokyo, Japan; TechnoSuruga Laboratory Co. Ltd., Shizuoka, Japan SHUICHI NAKAMURA • Department of Applied Physics, Graduate School of Engineering, Tohoku University, Sendai, Miyagi, Japan

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DAISUKE NAKANE • Department of Engineering Science, Graduate School of Informatics and Engineering, The University of Electro-Communications, Tokyo, Japan KEIICHI NAMBA • Graduate School of Frontier Biosciences, Osaka University, Suita, Osaka, Japan; RIKEN Center for Biosystems Dynamics Research, Suita, Osaka, Japan; RIKEN SPring-8 Center, Suita, Osaka, Japan; JEOL YOKOGUSHI Research Alliance Laboratories, Osaka University, Suita, Osaka, Japan TATSURO NISHIKINO • Institute for Protein Research, Osaka University, Suita, Osaka, Japan TAKAYUKI NISHIZAKA • Department of Physics, Gakushuin University, Tokyo, Japan LEON PALAO III • Department of Biochemistry and Biophysics, Perelman School of Medicine, University of Pennsylvania, Philadelphia, PA, USA JONATHAN D. PARTRIDGE • Department of Molecular Biosciences, The University of Texas at Austin, Austin, TX, USA GUOLIANG QIAN • College of Plant Protection, State Key Laboratory of Biological Interactions and Crop Health, Key Laboratory of Integrated Management of Crop Diseases and Pests, Nanjing Agricultural University, Nanjing, People’s Republic of China CHIKARA SATO • School of Integrative and Global Majors (SIGMA), University of Tsukuba, Tsukuba, Japan; Biological Science Course, Graduate School of Science and Engineering, Aoyama Gakuin University, Fuchinobe, Japan; Division of Immune Homeostasis, Department of Pathology and Microbiology, Nihon University School of Medicine, Itabashi, Tokyo, Japan; Division of Microbiology, Department of Pathology and Microbiology, Nihon University School of Medicine, Itabashi, Tokyo, Japan KEIKO SATO • Department of Frontier Oral Science, Medical and Dental Sciences, Graduate School of Biomedical Sciences, Nagasaki University, Sakamoto, Nagasaki, Japan XIAOLONG SHAO • College of Plant Protection, State Key Laboratory of Biological Interactions and Crop Health, Key Laboratory of Integrated Management of Crop Diseases and Pests, Nanjing Agricultural University, Nanjing, People’s Republic of China SATOSHI SHIBATA • Division of Bacteriology, Department of Microbiology and Immunology, Faculty of Medicine, Tottori University, Tottori, Japan; Department of Microbiology and Immunology, Faculty of Medicine, Tottori University, Tottori, Japan SHAMPHAVI SIVABALASARMA • Molecular Biology of Archaea, Faculty of Biology, Institute of Biology II, University of Freiburg, Freiburg, Germany; Spemann Graduate School of Biology and Medicine, University of Freiburg, Freiburg, Germany STEFAN STEIMLE • Department of Biochemistry and Biophysics, Perelman School of Medicine, University of Pennsylvania, Philadelphia, PA, USA MITSUHIRO SUGAWA • Graduate School of Arts & Sciences, The University of Tokyo, Tokyo, Japan YUHEI O. TAHARA • Graduate School of Science, Osaka City University, Osaka, Japan; Graduate School of Science, Osaka Metropolitan University, Osaka, Japan; The OCU Advanced Research Institute for Natural Science and Technology (OCARINA), Osaka City University, Osaka, Japan; The OMU Advanced Research Center for Natural Science and Technology, Osaka Metropolitan University, Osaka, Japan KYOSUKE TAKABE • Faculty of Life and Environmental Sciences, University of Tsukuba, Tsukuba, Ibaraki, Japan DAICHI TAKAHASHI • Graduate School of Science, Osaka City University, Osaka, Japan; Graduate School of Science, Osaka Metropolitan University, Osaka, Japan

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Contributors

AZUMA TAOKA • Institute of Science and Engineering, Kanazawa University, Kanazawa, Ishikawa, Japan NAOYA TERAHARA • Department of Physics, Chuo University, Tokyo, Japan HIROYUKI TERASHIMA • Department of Bacteriology, Institute of Tropical Medicine (NEKKEN), Nagasaki University, Nagasaki, Japan TAKUMA TOYONAGA • Graduate School of Science, Osaka City University, Osaka, Japan; Graduate School of Science, Osaka Metropolitan University, Osaka, Japan; The OCU Advanced Research Institute for Natural Science and Technology (OCARINA), Osaka City University, Osaka, Japan; The OMU Advanced Research Center for Natural Science and Technology, Osaka Metropolitan University, Osaka, Japan CHAO-KAI TSENG • Department of Physics, National Central University, Taoyuan, Taiwan BINGXIN WANG • College of Plant Protection, State Key Laboratory of Biological Interactions and Crop Health, Key Laboratory of Integrated Management of Crop Diseases and Pests, Nanjing Agricultural University, Nanjing, People’s Republic of China JUN XU • Department of Bacteriology, Graduate School of Medicine, University of the Ryukyus, Nishiharacho, Okinawa, Japan TOMOKO YAMAGUCHI • Graduate School of Frontier Biosciences, Osaka University, Suita, Osaka, Japan; RIKEN Center for Biosystems Dynamics Research, Suita, Osaka, Japan XIANG-YU ZHUANG • Department of Physics, National Central University, Taoyuan, Taiwan

Part I Bacterial Flagellar Protein Export and Assembly

Chapter 1 Purification of the Transmembrane Polypeptide Channel Complex of the Salmonella Flagellar Type III Secretion System Miki Kinoshita, Keiichi Namba, and Tohru Minamino Abstract Many motile bacteria employ the flagellar type III secretion system (fT3SS) to build the flagellum on the cell surface. The fT3SS consists of a transmembrane export gate complex, which acts as a proton/protein antiporter that couples proton flow with flagellar protein export, and a cytoplasmic ATPase ring complex, which works as an activator of the export gate complex. Three transmembrane proteins, FliP, FliQ, and FliR, form a core structure of the export gate complex, and this core complex serves as a polypeptide channel that allows flagellar structural subunits to be translocated across the cytoplasmic membrane. Here, we describe the methods for overproduction, solubilization, and purification of the Salmonella FliP/FliQ/ FliR complex. Key words Bacterial flagellum, Flagellar assembly, FliP, FliQ, FliR, Protein purification, Type III secretion system, Salmonella

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Introduction The flagellum of Salmonella enterica serovar Typhimurium (hereafter referred to as Salmonella) is a supramolecular motility machine consisting of the basal body, which acts as a bidirectional rotary motor powered by proton motive force (PMF) across the cytoplasmic membrane, the filament, which functions as a helical propeller to produce the thrust, and the hook, which exists between the basal body and filament and works as a universal joint to transmit torque produced by the motor to the filament [1, 2]. To construct the flagellum beyond the cytoplasmic membrane, flagellar structural subunits are transported via the flagellar type III secretion system (fT3SS) from the cytoplasm to the distal end of the growing flagellar structure. The fT3SS is composed a transmembrane export gate complex and a cytoplasmic ATPase ring complex (Fig. 1) [3–5].

Tohru Minamino et al. (eds.), Bacterial and Archaeal Motility, Methods in Molecular Biology, vol. 2646, https://doi.org/10.1007/978-1-0716-3060-0_1, © The Author(s), under exclusive license to Springer Science+Business Media, LLC, part of Springer Nature 2023

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Fig. 1 Schematic diagram of the flagellar type III protein export apparatus. The flagellar type III protein export apparatus consists of a transmembrane export gate complex made of FlhA, FlhB, FliP, FliQ, and FliR and a cytoplasmic ATPase ring complex consisting of FliH, FliI, and FliJ. The export gate complex is located within the central pore of the basal body MS-ring of the flagellum and acts as a proton/protein antiporter that couples proton flow with protein translocation across the cytoplasmic membrane. The export gate complex is activated by ATP hydrolysis by the ATPase ring complex. The ATPase ring complex associates with the basal body through the interaction of FliH with FliN in the C-ring. CM cytoplasmic membrane

The transmembrane export gate complex is located inside the basal body MS-ring formed by 34 copies of a transmembrane protein, FliF [6, 7], and is powered by PMF across the cytoplasmic membrane [8, 9]. Transmembrane proteins, FlhA, FlhB, FliP, FliQ, and FliR, assemble into the export gate complex with a stoichiometry of 9 FlhA, 1 FlhB, 5 FliP, 4 FliQ, and 1 FliR [10–13], and the export gate acts as a proton/protein antiporter that couples inward-directed proton flow with outward-directed protein translocation across the cytoplasmic membrane [14–16]. FliP, FliR, and FliQ form a polypeptide channel complex for the translocation of flagellar structural subunits across the cytoplasmic membrane (Fig. 2) [17, 18]. FlhA serves as an ion-driven export engine that conducts either protons or sodium ions and transports flagellar structural subunits through the polypeptide channel [19– 22]. The cytoplasmic ATPase ring complex is composed of 12 copies of FliH, 6 copies of FliI, and a single copy of FliJ [5]. The export gate complex requires ATP hydrolysis by the FliI ATPase to become a highly efficient proton/protein antiporter through an interaction between FlhA and FliJ [14, 16, 20, 22]. FlhB associates with the FliP5-FliQ4-FliR1 complex and plays an important role in the

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Fig. 2 CryoEM structure of the FliP5-FliQ4-FliR complex. Cα ribbon representation of the FliP5-FliQ4-FliR1 complex (PDB ID: 6R69). Green, FliP; blue, FliQ; yellow, FliR. FliP and FliR require the FliO scaffold protein to assemble into the FliP5-FliR1 complex in the cytoplasmic membrane. Four copies of FliQ associate with the outside of the FliP5-FliR1 complex. The central pore of the FliP5-FliR1 complex serves as a polypeptide channel for the translocation of flagellar structural subunits across the cytoplasmic membrane

gating mechanism of the fT3SS along with FlhA and the cytoplasmic ATPase complex [13, 16, 23]. The assembly of the transmembrane export gate complex begins with formation of the FliP5-FliR1 complex with a help of the FliO scaffold protein [17, 18]. Then, four FliQ molecules are associated with the FliP5-FliR1 complex on its outside (Fig. 2), followed by the assembly of FlhB through interactions of FlhB with FliP, FliQ, and FliR [11, 13]. Finally, nine FlhA subunits bind to the FliP5-FliQ4-FliR1-FlhB1 complex during MS-ring formation [24]. This book chapter describes the protocols we used for overexpression, solubilization, and purification of the Salmonella FliP5FliQ4FliR1 complex.

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Materials Prepare all solutions using ultrapure water and analytical grade reagents, and then autoclave them at 121  C for 20 min.

2.1 Salmonella Strains and Plasmids

1. JR501 (see Note 1) [25]. 2. SJW1368 Δ(cheW-flhD) (see Note 2) [26]. 3. pKY077 (pTrc99AFF4/FliO + His-FliP + HA-FliQ + FliRFLAG) (see Note 3) [18].

2.2

Culture Media

1. Luria broth (LB): 1.0% (w/v) Bacto tryptone, 0.5% (w/v) Yeast extract, 0.5% (w/v) NaCl.

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2. LB agar (LA) plate: 1.0% (w/v) Bacto tryptone, 0.5% (w/v) Yeast extract, 0.5% (w/v) NaCl, 1.5% (w/v) Bacto agar. 3. 2YT: 1.6% (w/v) Bacto tryptone, 1.0% (w/v) Yeast extract, 0.5% (w/v) NaCl. 4. Ampicillin. 2.3 Cell Growth and Harvest

1. 37  C static incubator. 2. 16  C, 30  C, and 37  C shakers. 3. Spectrophotometer. 4. Disposable cuvette. 5. Test tubes. 6. Erlenmeyer flask (100 mL, 5.0 L). 7. Floor-standing centrifuge with rotors (e.g., Avanti J-E, JA-20 rotor, JLA-10.500 rotor, Beckman Coulter). 8. Centrifuge tubes (50 mL, 500 mL). 9. Microtubes (5.0 mL, 1.5 mL). 10.

2.4

Transformation

80  C freezer.

1. 0.1 M MgCl2. Store at 4  C. 2. 0.1 M CaCl2. Store at 4  C. 3. 50%(w/v) glycerol. Store at 4  C. 4. Floor-standing centrifuge with rotors. 5. Centrifuge tubes (50 mL). 6. Refrigerated microcentrifuge with rotors. 7. 1.5 mL microtubes. 8. 42  C heating block. 9. 37  C incubator. 10. Ice bucket. 11. Ice.

2.5 Preparation and Solubilization of Cellular Membranes

1. Lysis buffer: 20 mM Tris–HCl, pH 8.0, 3.0 mM EDTA. Store at 4  C. 2. Membrane fractionation buffer: 50 mM Tris–HCl, pH 8.0, 300 mM NaCl, 10% (w/v) glycerol. Store at 4  C. 3. 2x Solubilization buffer: 50 mM Tris–HCl, pH 8.0, 300 mM NaCl, 40 mM imidazole. Store at 4  C. 4. Lauryl maltose neopentyl glycol (LMNG). 5. Ultrasonic disintegrator. 6. 100 mL stainless steel beaker. 7. Homogenizer.

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8. Floor-standing centrifuge with rotors. 9. Centrifuge tubes. 10. Floor-standing ultracentrifuge with rotors (e.g., Optima XE-90, Beckman Coulter). 11. Desktop ultracentrifuge with rotors (e.g., Optima MAX-TL, Beckman Coulter). 12. Ultracentrifuge tubes. 13. Refrigerated microcentrifuge with rotors. 14. 5.0 mL microtubes. 15. Ice bucket. 16. Ice. 2.6 Measurement of Protein Concentration in Crude Membrane Fractions

1. A buffer: 2.0%(w/v) Na2CO3, 0.1 N NaOH. Store at 4  C. 2. B1 buffer: 0.5%(w/v) CuSO4•5H2O. Store at 4  C. 3. B2 buffer: 1.0% (w/v) sodium citrate. Store at 4  C. 4. Freshly prepared C buffer: 50 volumes of A buffer, 1 volume of B1 buffer, 1 volume of B2 buffer. 5. Folin & Ciocalteu’s phenol reagent. 6. 10 mg/mL bovine serum albumin (BSA). 7. 1.5 mL microtubes. 8. Spectrophotometer. 9. Disposable cuvette.

2.7 Purification of the FliP/FliQ/FliR Complex

1. 2 Mix buffer: 100 mM Tris–HCl, pH 8.0, 600 mM NaCl, 10% (w/v) glycerol. Store at 4  C. 2. 2 M imidazole-HCl, pH 8.0. Store at 4  C. 3. 1.0% (w/v) LMNG 4. Cold ultrapure water. 5. SEC buffer: 20 mM Tris–HCl, pH 8.0, 300 mM NaCl, 2.0 mM EDTA, 5.0% (w/v) glycerol, 0.005% LMNG. 6. 5.0 mL polypropylene columns. 7. Ni2+-nitrilotriacetic acid (NTA) resin. 8. Tube rotator. 9. Size exclusion chromatography (SEC) column [e.g., Superdex 200 10/300 (GE Healthcare)]. 10. Fast protein liquid chromatography (FPLC) system. 11. Chromato chamber. 12. Disposable test tubes. 13. 10 mL high-clarity polypropylene conical centrifugation tubes. 14. 1.5 mL microtubes.

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2.8 Identification of Protein Expression and Purified Protein Sample

1. 2 SDS loading buffer: 125 mM Tris–HCl, pH 6.8, 4% (w/v) sodium dodecyl sulfate (SDS), 20% (w/v) glycerol, 0.002% (w/v) bromophenol blue. 2. 2-mercaptoethanol. 3. 95  C heating block. 4. Plastic containers. 5. Electrophoresis apparatus for sodium dodecyl polyacrylamide gel electrophoresis (SDS-PAGE).

sulfate-

6. 12.5% SDS-polyacrylamide gels. 7. Tris-Tricine running buffer for SDS-PAGE: 12.1 g Tris (hydroxymethyl)aminomethane, 17.9 g tricine, 1.0 g SDS per liter. 8. Coomassie Brilliant blue (CBB) staining solution: One Coomassie tablet in 1 L 20% (v/v) ethanol, 10% (v/v) glacial acetic acid. 9. Distaining solution: 10% (v/v) ethanol, 7% (v/v) glacial acetic acid. 10. Nitrocellulose membranes. 11. Blotting apparatus. 12. Transfer buffer: 25 mM, Tris(hydroxymethyl)aminomethane, 250 mM glycine, 20% (v/v) methanol. 13. TBS-T: 20 mM Tris–HCl, pH 7.5, 500 mM NaCl, 0.1% (v/v) Tween-20. 14. Blocking buffer: 5% skim milk in TBS-T. 15. Polyclonal antibodies against FliO and FliP were produced by MBL (Nagoya, Japan). 16. Monoclonal anti-HA antibody. 17. Monoclonal anti-FLAG antibody. 18. Goat anti-rabbit IgG-HRP. 19. Goat anti-mouse IgG-HRP. 20. Chemiluminescence reagents (e.g., ECL Prime immunoblotting detection kit). 21. Chemiluminescence detection system. 2.9 Observation of Purified FliP/FliQ/FliR Complex by Electron Microscopy

1. Glass slide. 2. Tweezer. 3. Ion coater. 4. Desiccator. 5. Paper filter. 6. Parafilm.

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7. Carbon-coated copper grids. 8. 2.0% (w/v) uranyl acetate. 9. Transmission electron microscope.

3 3.1

Methods Transformation

1. Inoculate a single colony of the Salmonella SJW1368 strain into a 5.0 mL volume of LB in a test tube. 2. Incubate the culture overnight at 30  C with shaking. 3. Add 0.3 mL of the overnight culture to a 30 mL volume of fresh LB in a 100 mL Erlenmeyer flask. 4. Measure OD600 of the culture using a spectrophotometer. 5. Incubate the culture at 37  C with shaking until the cell density reaches an OD600 of 0.6–0.8. 6. Transfer the culture into a 50 mL centrifuge tube. 7. Centrifuge the tube (8000 g, 5 min, 4  C). 8. Discard the supernatant and suspend the cell pellet in 30 mL of cold 0.1 M MgCl2. 9. Collect the cells by centrifugation (8000 g, 5 min, 4  C), and suspend the harvested cells in 15 mL of cold 0.1 M CaCl2. 10. Leave the tube on ice for 30 min. 11. Collect the cells by centrifugation (8000 g, 5 min, 4  C), and suspend the harvested cells in 1.0 mL of cold 0.1 M CaCl2 and 1.0 mL of cold 50% (w/v) glycerol. 12. Competent cells are either used immediately or stored at 80  C. 13. Add a 125 μL volume of the competent cells into a 1.5 mL microtube containing 1–2 μL of the pKY077 plasmid prepared from the Salmonella JR501 strain. 14. Leave the tube on ice for 30 min. 15. Heat the tube at 42  C for 2 min. 16. Leave the tube on ice for 5 min. 17. Add fresh 1.0 mL of LB to the tube. 18. Incubate the tube at 37  C for 1 h. 19. Collect the cells by centrifugation (9000 g, 5 min, 4  C), and resuspend the harvested cells in 100 μL of LB. 20. Streak the cell suspension to an LA plate containing 50 μg/mL ampicillin. 21. Leave the plate overnight in a 37  C incubator.

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3.2 Cell Culture and Harvest

1. Inoculate a single fresh transformant into a 30 mL volume of 2YT containing 100 μg/mL ampicillin. 2. Proliferate cells overnight at 30  C with shaking. 3. Add 13 mL of the overnight culture to a 1.3 L volume of fresh 2YT in a 5.0 L Erlenmeyer flask. 4. Measure OD600 of each culture using a spectrophotometer. 5. Incubate the culture at 30  C with shaking until the cell density reaches an OD600 of 0.6. 6. Leave the culture at 4  C for 30 min. 7. Incubate the culture at 16  C for 24 h with shaking. 8. Transfer the culture into a 500 mL centrifuge tube. 9. Collect the cells by centrifugation (12,000 g, 5 min, 4  C), and store the harvested cells at 80  C until the membrane fraction is prepared.

3.3 Preparation of Membrane Fractions

1. Suspend cells, which have been harvested from the 1.3 L culture (Subheading 3.2), in a 50 mL volume of cold Lysis buffer. 2. Transfer the cell suspension to a 100 mL stainless steel beaker. 3. Keep the beaker on ice. 4. Disrupt the cells by sonication for 10 min. 5. Remove cell debris and undisrupted cells by centrifugation (20,000 g, 10 min, 4  C). 6. Harvest a crude membrane fraction by ultracentrifugation (110,000 g, 1 h, 4  C). 7. Suspend the membrane fraction in a 5.0 mL volume of cold membrane fraction buffer using a homogenizer. 8. Measure the protein concentration of the membrane fraction by Lowry method (Subheading 3.4) [27]. 9. Adjust the protein concentration to 20 mg/mL with cold membrane fraction buffer. 10. Divide 5.0 mL each into 5.0 mL microtubes, and store at 80  C until use.

3.4 Measurement of Protein Concentration by the Lowry Method

1. Dilute membrane fraction (Subheading 3.3) 100-fold with cold membrane fraction buffer. 2. Mix 100 μL of diluted membrane fraction and 1.0 mL freshly prepared C buffer in a 1.5 mL microtube and voltex. 3. Leave the tube for 10 min at room temperature. 4. Add 100 μL of Folin & Ciocalteu’s phenol reagent to the tube and voltex well. 5. Leave the tube for 3 min at room temperature.

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6. Measure of OD750 using a spectrophotometer. 7. Create a standard curve of BSA with various concentrations. 3.5 Solubilization of Membrane Fractions by LMNG

1. Thaw 5.0 mL frozen crude membrane fractions on ice. 2. Transfer 5.0 mL of the membrane fractions to a 30 mL beaker, and then add 5.0 mL of fleshly prepared solubilization buffer (2.5 mL of 2 solubilization buffer, 2.5 mL of ultrapure water) containing 2.0%(w/v) LMNG. 3. Gently stir the mixture at 4  C for 1 h. 4. Collect the supernatant by ultracentrifugation (110,000 g, 4  C, 1 h).

3.6 Purification of the FliP/FliQ/FliR Complex

1. Add a 1.0 mL volume of the Ni-NTA agarose resin to a 10 mL high-clarity polypropylene conical centrifuge tube, and equilibrate the resin with a 15 mL volume of freshly prepared Wash buffer-20 (7.5 mL of 2 Mix buffer, 150 μL of 2 M imidazole, 150 μL of 1.0%(w/v) LMNG, 7.2 mL of ultrapure water). 2. Transfer the supernatants (subheading 3.5, step 1) into the tube. 3. Mix well at 4  C for 1 h using a tube rotator. 4. Pack the resin into a 5.0 ml polypropylene column. 5. Wash the column with a 15 mL volume of freshly prepared Wash buffer-20. 6. Wash the column with a 2.0 mL volume of freshly prepared Wash buffer-50 (1.0 mL of 2 Mix buffer, 50 μL of 2 M imidazole, 20 μL of 1.0%(w/v) LMNG, 0.93 mL of ultrapure water). 7. Wash the column with a 2.0 mL volume of freshly prepared Wash buffer-80 (1.0 mL of 2 Mix buffer, 80 μL of 2 M imidazole, 20 μL of 1.0%(w/v) LMNG, 0.9 mL of ultrapure water). 8. Elute bound proteins with a 2.0 mL volume of freshly prepared Elution buffer-100 (1.0 mL of 2 Mix buffer, 100 μL of 2 M imidazole, 20 μL of 1.0%(w/v) LMNG, 0.88 mL of ultrapure water). 9. Elute bound proteins with a 5.0 mL volume of freshly prepared Elution buffer-400 (2.5 mL of 2 Mix buffer, 1.0 mL of 2 M imidazole, 50 μL of 1.0%(w/v) LMNG, 1.45 mL of ultrapure water). 10. Run proteins in each fraction on SDS-PAGE, followed by CBB staining and immunoblotting to identify fractions containing FliO, His-FliP, HA-FliQ, and FliR-FLAG (see Note 4). 11. Collect and pool fractions containing FliO, His-FliP, HA-FliQ, and FliR-FLAG.

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Fig. 3 Purification of the His-FliP/HA-FliQ/FliR-FLAG complex by size exclusion chromatography. (a) Elution profile of the His-FliP/HA-FliQ/FliR-FRAG complex from a Superdex 200 10/300 column equilibrated with 20 mM Tris–HCl, pH 8.0, 300 mM NaCl, 2 mM EDTA, 5% glycerol, 0.005% (w/v) LMNG. Membrane fractions were solubilized by 1.0% (w/v) LMNG. Then, the protein complex was purified by Ni affinity chromatography, followed by size exclusion chromatography. The peak 1 contains the FliO/His-FliP/HA-FliQ/FliR-FLAG as judged by immunoblotting, whereas the peak 2 contains the His-FliP/HA-FliQ/FliR-FLAG complex. (b) Electron micrograph of negatively stained His-FliP/HA-FliQ/FliR-FLAG complex

12. Load 500 μL of pooled fractions onto a SEC column (e.g., Superdex 200 10/300) pre-equilibrated with cold SEC buffer. 13. Pass proteins over the SEC column using FPLC at the flow rate of 0.4 mL/min at 4  C. 14. Collect eluted fractions using a fraction collector (Fig. 3a). 15. Run proteins in each fraction on SDS-PAGE, followed by CBB staining and immunoblotting to identify fractions containing the His-FliP/HA-FliQ/FliR-FLAG complex (see Note 5). 3.7 Observation of Negatively Stained FliP/FliQ/FliR Complex by Electron Microscopy

1. Glow-charge carbon-coated copper grids on a grass slide for 20 s using an ion motor operating at a constant current of 20 mA. 2. Apply 3.0 μL of purified His-FliP/HA-FliQ/FliR-FLAG complex onto a carbon-coated copper grid. 3. Remove the extra solution using a paper filter. 4. Place 3.0 μL of 2.0% (w/v) uranyl acetate on a parafilm. 5. Put the sample glid on the droplet of 2.0% (w/v) uranyl acetate on a parafilm. 6. Remove the extra solution using a paper filter. 7. Repeat steps 4–6 twice.

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8. Place negatively stained grid on a filter paper and dry it in desiccator at least for 30 min. 9. Observe negatively stained samples by a transmission electron microscope operating at 100 kV (Fig. 3b).

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Notes 1. The Salmonella JR501 strain is used for conversion of E. coliderived plasmids to compatibility with Salmonella [24]. 2. The Salmonella SJW1368 strain does not have master regulatory genes, and neither flagellar genes, motor genes nor chemotaxis genes are transcribed [25]. 3. Because FliP has a cleavable signal peptide at its N-terminus, the His tag (LHHHHHH) is inserted between Gln-22 and Leu-23 of FliP for rapid and efficient purification [18]. To identify FliQ and FliR by immunoblotting, the HA tag (YPYDVPDYA) and the FLAG tag (DYKDDDDK) are fused to the N-terminus of FliQ and the C-terminus of FliR, respectively [18]. Because FliP requires FliO to efficiently form oligomer in the membrane, the pKY077 plasmid also encodes FliO along with His-FliP, HA-FliQ, and FliR-FRAG [17, 18]. 4. A 10 μL volume of each fraction, a 10 μL volume of 2 SDS loading buffer, and a 1 μL volume of 2-mercaptoethanol are mixed, followed by incubation at 95  C for 3 min. A protein standard maker and purified protein samples are loaded to each lane, and electrophorese is performed in the Tris/Tricin buffer at a constant voltage of 125 V until the sample fractions reach the bottom of the gel. Following the electrophoresis, proteins in the gel are fixed and stained by CBB. To carry out immunoblotting, proteins are transferred onto a nitrocellulose membrane in the transfer buffer using a blotting apparatus. The membrane is transferred into a 25 mL volume of the Blocking buffer and is gently shaken for 1 h at room temperature. After washing one with 25 mL of TBS-T, the membrane is incubated in 10 mL of TBS-T containing polyclonal anti-FliO, polyclonal anti-FliP, monoclonal anti-HA, or monoclonal anti-FLAG antibody (normally 1:10,000 dilution) for 1 h at room temperature with gently shaking. After washing three times with 25 mL of TBS-T, the membrane is incubated in 10 mL of TBS-T containing goat anti-rabbit IgG-HRP or goat antirabbit IgG-HRP (1:2500 dilution) for 1 h at room temperature with gently shaking. Immunodetection is performed using chemiluminescence reagents and chemiluminescence detection system.

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5. It has been shown that the FliO complex easily dissociates from the FliP/FliQ/FliR complex during storage of purified FliO/ FliP/FliQ/FliR complex at 4  C for several days [18]. So, we always run the Peak 1 fractions again on the SEC column to purify the FliO complex and the FliP/FliQ/FliR complex separately.

Acknowledgments This work was supported in part by JSPS KAKENHI Grant Numbers JP20K15749 and JP22K06162 (to M.K.) and JP19H03182, JP22H02573, and JP22K19274 (to T.M.) and MEXT KAKENHI Grant Number JP20H05532 and JP22H04844 (to T.M.). This work has also been supported by Platform Project for Supporting Drug Discovery and Life Science Research (BINDS) from AMED under Grant Number JP19am0101117 to K.N., by the Cyclic Innovation for Clinical Empowerment (CiCLE) from AMED under Grant Number JP17pc0101020 to K.N. and by JEOL YOKOGUSHI Research Alliance Laboratories of Osaka University to K.N. References 1. Morimoto YV, Minamino T (2014) Structure and function of the bi-directional bacterial flagellar motor. Biomol Ther 4:217–234 2. Nakamura S, Minamino T (2019) Flagelladriven motility of bacteria. Biomolecules 9:279 3. Minamino T (2014) Protein export through the bacterial flagellar type III export pathway. Biochim Biophys Acta 1843:1642–1648 4. Minamino T (2018) Hierarchical protein export mechanism of the bacterial flagellar type III protein export apparatus. FEMS Microbiol Lett 365:fny117 5. Minamino T, Kawamoto A, Kinoshita M et al (2020) Molecular organization and assembly of the export apparatus of flagellar type III secretion systems. Curr Top Microbiol Immunol 427:91–107 6. Johnson S, Furlong EJ, Deme JC et al (2021) Molecular structure of the intact bacterial flagellar basal body. Nat Microbiol 6:712–721 7. Kawamoto A, Miyata T, Makino F et al (2021) Native flagellar MS ring is formed by 34 subunits with 23-fold and 11-fold subsymmetries. Nat Commun 12:4223 8. Minamino T, Namba K (2008) Distinct roles of the FliI ATPase and proton motive force in bacterial flagellar protein export. Nature 451: 485–488

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Purification of the Salmonella FliPQR Complex 16. Minamino T, Morimoto YV, Kinoshita M et al (2021) Membrane voltage-dependent activation mechanism of the bacterial flagellar protein export apparatus. Proc Natl Acad Sci U S A 118:e2026587118 17. Fabiani FD, Renault TT, Peters B et al (2017) A flagellum-specific chaperone facilitates assembly of the core type III export apparatus of the bacterial flagellum. PLoS Biol 15: e2002267 18. Fukumura T, Makino F, Dietsche T et al (2017) Assembly and stoichiometry of the core structure of the bacterial flagellar type III export gate complex. PLoS Biol 15:e2002281 19. Hara N, Namba K, Minamino T (2011) Genetic characterization of conserved charged residues in the bacterial flagellar type III export protein FlhA. PLoS One 6:e22417 20. Minamino T, Morimoto YV, Hara N et al (2016) The bacterial flagellar type III export gate complex is a dual fuel engine that can use both H+ and Na+ for flagellar protein export. PLoS Pathog 12:e1005495 21. Erhardt M, Wheatley P, Kim EA et al (2017) Mechanism of type-III protein secretion: regulation of FlhA conformation by a functionally

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critical charged-residue cluster. Mol Microbiol 104:234–249 22. Minamino T, Kinoshita M, Morimoto YV et al (2021) The FlgN chaperone activates the Na+driven engine of the Salmonella flagellar protein export apparatus. Commun Biol 4:335 23. Kinoshita M, Namba K, Minamino T (2021) A positive charge region of Salmonella FliI is required for ATPase formation and efficient flagellar protein export. Commun Biol 4:464 24. Morimoto YV, Ito M, Hiraoka KD, Che Y-S, Bai F, Kami-ike N, Namba K, Minamino T (2014) Assembly and stoichiometry of FliF and FlhA in Salmonella flagellar basal body. Mol Microbiol 91:1214–1226 25. Ryu J, Hartin RJ (1990) Quick transformation in Salmonella typhimurium LT2. BioTechniques 8:43–45 26. Ohnishi K, Ohto Y, Aizawa S, Macnab RM, Iino T (1994) FlgD is a scaffolding protein needed for flagellar hook assembly in Salmonella typhimurium. J Bacteriol 176:2272– 2281 27. Lowry OH, Rosebrough NJ, Farr AL, Randall RJ (1951) Protein measurement with the Folin phenol reagent. J Biol Chem 193:265–275

Chapter 2 In Vitro Flagellar Type III Protein Transport Assay Using Inverted Membrane Vesicles Katsumi Imada and Hiroyuki Terashima Abstract The flagellar axial proteins are transported across the cytoplasmic membrane into the central channel of the growing flagellum via the flagellar protein export apparatus, a member of the type III secretion system (T3SS). To reveal the molecular mechanism of protein transport by the T3SS, accurate measurement of protein transport under various conditions is essential. In this chapter, we describe an in vitro method for flagellar protein transport assay using inverted membrane vesicles (IMVs) prepared from Salmonella cells. This method can easily and precisely control the condition around the T3SS and be applied to other T3SSs. Key words T3SS, Protein transport assay, Inverted membrane vesicle

1 1.1

Introduction Background

The bacterial flagellum is a long filamentous organelle for motility and consists of a tubular axial structure and basal body rings. The axial structure extends outward from the MS ring of the basal body embedded in the cell membrane. The axial structure of Salmonella typhimurium is made up of more than 20,000 protein subunits of 10 different proteins. These proteins are translocated via the flagellar protein export apparatus across the cytoplasmic membrane into the central channel of the growing flagellum [1, 2] (Fig. 1a). The flagellar protein export apparatus is a member of the type III secretion system (T3SS) and thus is called fT3SS. The fT3SS consists of the export gate formed by FlhA, FlhB, FliP, FliQ, and FliR within the transmembrane MS ring of the basal body and the cytoplasmic ATPase complex composed of FliH, FliI, and FliJ [3– 5]. FliH and FliI also form the FliH2/FliI complex in the cytoplasm and interact with export substrates and export chaperones [6–8]. Many genetic and biochemical studies have revealed that efficient protein export by fT3SS is driven by proton motive force (PMF) and ATP-hydrolysis by the cytoplasmic ATPase complex

Tohru Minamino et al. (eds.), Bacterial and Archaeal Motility, Methods in Molecular Biology, vol. 2646, https://doi.org/10.1007/978-1-0716-3060-0_2, © The Author(s), under exclusive license to Springer Science+Business Media, LLC, part of Springer Nature 2023

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Fig. 1 Schematic diagram of the Salmonella flagellum (a) and the in vitro transport assay method using the inverted membrane vesicle (IMV) (b). (a) The filamentous part consisting of the rod, the hook, the hookfilament junction, and the filament are called the flagellar axial structure. The fT3SS consists of the export gate and the cytoplasmic ATPase complex. CM, the cytoplasmic membrane; PG, the peptidoglycan layer; OM, the outer membrane. (b) The IMV was filled with 300 mM NaCl at pH 6.0 and suspended in solution with 125 mM K+ and 5 mM MgCl2 at pH 7.5. Endogenous FoF1-ATP synthase transfers proton into the IMV using ATP-hydrolysis energy

[9–13]. However, the molecular mechanism of protein export is still not well understood. To fully understand the protein transport mechanism, quantitative measurement of protein transport under various conditions is essential, but control of the measurement conditions around T3SS is very difficult in vivo. Therefore, an in vitro transport assay method that enables precise control of the conditions is required. Small transporters can be reconstructed in liposomes. However, since the fT3SS is a huge molecular assembly, it is difficult to purify and reconstruct functional T3SS in a lipid bilayer or a liposome. Inverted membrane vesicle (IMV) has been used for studying protein translocators, such as Sec translocon [14] and twin arginine translocator [15]. Thus, we focused on IMV. We recently developed the IMV method, applied it to the Salmonella fT3SS, and succeeded to measure flagellar protein transport under various conditions [16–18] (Fig. 1b). Here we describe the details of our method. This IMV method can be applied to other T3SS, if the cytoplasmic T3SS component proteins can be purified. 1.2 Overview of the Method

IMVs are prepared from spheroplasts of Salmonella cells. The cells are converted to spheroplasts by incubation with lysozyme and EDTA to remove the peptidoglycan layer and the outer membrane, respectively. The spheroplasts are disrupted by a high-pressure

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homogenizer to produce IMVs. The IMVs are purified by sucrose density gradient centrifugation and can be stored in a deep freezer at 80 . The IMV solution is filtered, and the IMVs are further purified with a Sephadex column right before the assay. The protein export reaction is initiated by adding ATP (Fig. 1b). PMF is generated by the reverse reaction of endogenous FoF1-ATP synthase embedded in the IMVs. The external solution needs to contain the cytoplasmic ATPase component proteins and substrate proteins. Therefore, these proteins should be purified before the assay. The purification methods have been described previously [16]. The protein export is terminated by the addition of Proteinase K, which degrades the remaining substrates in the assay mixture as well as the cytoplasmic ATPase component proteins and the cytoplasmic domains of the export apparatus. The proteins transported into the IMVs are analyzed by immunoblotting with an antibody against the export substrate. FlgD, the hook cap protein, is used as an export substrate in this protocol.

2

Materials Prepare all solution using Milli-Q water, and use analytical grade reagents.

2.1 Salmonella enterica Strain and Plasmid 2.2

Culture

1. STH001 (ΔflhB ΔflgD ΔfliT): A Salmonella mutant strain with deletion of the flhB, flgD, and fliT genes [16]. 2. pITH104: pBAD33SD-flhB(N269A) + flhDC [16]. 1. Luria broth (LB): 1% (w/v) Bacto tryptone, 0.5% (w/v) Yeast extract, 0.5% (w/v) NaCl. Autoclave at 121  C for 20 min. 2. LB agar plate (LA): 1% (w/v) Bacto tryptone, 0.5% (w/v) Yeast extract, 0.5% (w/v) NaCl, 1.5% (w/v) Bacto agar. Autoclave at 121  C for 20 min. 3. 50 mg/mL Ampicillin solution. 4. 30 mg/mL Chloramphenicol solution. 5. 20% (w/v) L-arabinose solution. 6. Incubator. 7. Shaker.

2.3 Preparation of IMV Stock

1. High-speed centrifuge. 2. Sucrose solution: 10 mM Tris–HCl pH 8.0, 0.75 M sucrose. 3. Lysozyme. 4. 1.5 mM EDTA solution. 5. Magnetic stirrer. 6. Dark field microscope.

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2.4 Preparation of IMV

1. High-speed refrigerated centrifuge. 2. Solution A: 20 mM MES-NaOH pH 6.0, 300 mM NaCl (see Note 1). 3. Protease inhibitor cocktail (e.g., cOmplete EDTA-free, Roche). 4. Pressure cell homogenizer (e.g., SPCH10, STANSTED). 5. Ultracentrifuge tube (e.g., 25  64 mm, 27 mL, Beckman Coulter). 6. Ultracentrifuge. 7. 60% (w/w) sucrose: 24 g sucrose in 16 g of solution A. 8. 40% (w/w) sucrose: 18 g sucrose in 27 g of solution A. 9. 50% (w/w) sucrose: mix 10 mL of 60% sucrose with 10 mL of 40% sucrose. 10. 45% (w/w) sucrose: mix 5 mL of 60% sucrose with 15 mL of 40% sucrose. 11. Open-top thin polyclear tube (e.g., 25  89 mm, 38.5 mL, Seton). 12. Liquid nitrogen.

2.5 In Vitro Transport Assay

1. Polycarbonate membranes 0.8 μm 19 mm (e.g., Avanti polar lipids). 2. Mini-Extruder (e.g., Avanti polar lipids). 3. Sephadex G-50 fine resin (e.g., Cytiva). 4. Solution B: 125 mM KCl, 20 mM Tris–HCl pH 7.5. 5. Disposable plastic gravity flow column. 6. Spectrometer. 7. External solution: 20 mM Tris–HCl, pH 7.5, 125 mM KCl, 5 mM MgCl2, 1 mM dithiothreitol (DTT), 4 μM FlgD (substrate protein), 0.25 μM FliJ, and the 1.5 μM FliH2/FliI complex (The indicated concentrations are the final concentrations of the assay mixture.) (see Note 2). 8. 1.5 mL polyallomer tube with cap, for micro ultracentrifuge (e.g., Beckman Coulter). 9. Tabletop ultracentrifuge. 10. 0.1 M ATP solution. The ATP solution is prepared by dissolving ATP (dipotassium salt) in Tris solution (final concentration of 20 mM) followed by neutralization with KOH. 11. Heat block. 12. Proteinase K solution: Add 1 mg of proteinase K in 1 mL of solution B. 13. 1% (v/v) Triton X100 solution.

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14. Trichloroacetate (TCA). 15. High-speed refrigerated centrifuge. 16. SDS-PAGE loading buffer (2): 125 mM Tris–HCl, pH 6.8, 4% (w/v) sodium dodecyl sulfate (SDS), 20% (w/v) glycerol, 0.002% (w/v) bromophenol blue. 17. 15% SDS-polyacrylamide gel. 18. SDS-PAGE apparatus. 19. Tris-glycine buffer (SDS-PAGE running buffer): 30 g Tris, 144 g glycine, 10 g SDS in 1 L of milli-Q water. 20. Nitrocellulose membrane. 21. Immunoblotting apparatus. 22. Immunoblotting transfer buffer: 25 mM Tris, 250 mM glycine, 20% (v/v) methanol. 23. TBS containing Tween-20 (TBS-T): 20 mM Tris–HCl, pH 7.5, 500 mM NaCl, 0.1% (v/v) Tween-20. 24. Immunoblotting blocking buffer: 5% (w/v) skim milk in TBS-T. 25. Polyclonal antibodies against the substrate and FlhA. 26. Goat anti-rabbit IgG-HRP. 27. Chemiluminescence detection kit for immunoblot. 28. Chemiluminescence detection imaging system.

3 3.1

Methods Cell Culture

1. Streak the Salmonella cells strain STH001 (ΔflhB ΔflgD ΔfliT) harboring pITH104 on a flesh LB agar plate, and incubate the plate at 37  C overnight (see Note 3). 2. Inoculate a single colony from the plate into 40 mL of LB, and incubate the cells at 37  C for 8 h with shaking. 3. Inoculate 10 mL of the culture into 1 L of LB in 5 L flask, and incubate it at 30  C with shaking for 1 h. 4. Add 1 mL of 20% L-arabinose solution [final concentration of L-arabinose: 0.02% (w/v)], and continue the culture at 18  C for 12–16 h with shaking.

3.2 Preparation of Spheroplast

1. If OD600 will reach around 1.5, harvest cells by centrifugation (6700  g, 5 min, 4  C) using a fixed angle rotor (e.g., JLA10.500, Beckman Coulter). 2. Discard the supernatant (see Note 4).

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3. Add 75 mL of sucrose solution to the cell pellets, and suspend the pellet quickly by pipetting. Transfer the suspension into a beaker. 4. Add 22.5 mg of lysozyme on ice (final concentration: 0.1 mg/ mL), and stir the solution using a magnetic stirrer to completely dissolve lysozyme (typically 1–2 min). 5. Add 150 mL of 1.5 mM EDTA solution very slowly (over 5–8 min) on ice. 6. Gently stir the solution using a magnetic stirrer at 4  C for 1 h to produce spheroplasts. 7. Check spheroplast formation. Drop ~2 μL of the solution on a slide glass, and observe spheroplast by using a dark field microscope. 3.3 Preparation of IMV Stock

1. Collect spheroplasts by centrifugation (4600  g for 10 min, 4  C) using a fixed angle rotor (e.g., JLA10.500, Beckman Coulter). 2. Discard the supernatant (see Note 5). 3. Suspend the spheroplasts with 25 mL of solution A with 1/8 tablet of protease inhibitor cocktail (e.g., cOmplete EDTAfree, Roche) (see Note 6). 4. Load the suspension to a high-pressure cell homogenizer (STANSTED) at 70 MPa to produce IMV (see Note 7). 5. Collect IMV in a 50 mL plastic tube. 6. Spin cell lysates (15,300  g, 10 min, 4  C) using a fixed angle rotor (e.g., AR510-04 , TOMY) to remove debris. 7. Transfer the supernatant to a 27 mL ultracentrifuge tube, and fill the tube by adding solution A. 8. Spin the tube (213,000  g, 1 h, 4  C) using a fixed angle rotor (e.g., 50.2 Ti, Beckman Coulter) to precipitate IMVs. 9. Discard the supernatant. 10. Suspend the IMV pellets with 1.5 mL of solution A using a pipette. 11. Preparation of a sucrose density-gradient. Add 5 mL of 60% (w/w) sucrose, 9 mL of 50% (w/w) sucrose, 9 mL of 45% (w/w) sucrose, and 6 mL of 40% (w/w) sucrose in an open-top thin polyclear tube (25  89 mm) in this order. 12. Load the IMV solution on the sucrose density gradient in the polyclear tube. 13. Spin the tube (90,000  g, 16 h, 4  C) using a swing rotor (e.g., SW32 Ti, Beckman Coulter) with slow acceleration and deceleration.

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14. Collect the brown-colored layer (~2 mL), which contains IMVs, using a pipette. 15. Pour the IMV solution into an ultracentrifuge tube (27 mL), and add solution A to fill the tube. 16. Spin the tube (213,000  g, 1 h, 4  C) using a swing rotor (e.g., SW32 Ti, Beckman Coulter) to precipitate IMVs. 17. Discard the supernatant. 18. Suspend the IMV pellets with 1 mL of solution A using a pipette and a disposable homogenizer. 19. Divide the suspension into 300 μL aliquots, freeze them by liquid nitrogen, and store them in 80  C until before use. 3.4 Preparation of the IMV Solution for Transport Assay

1. Thaw a frozen stock of the IMV solution at room temperature. 2. Homogenize the solution through 0.8 μm polycarbonate membrane filter with the Avanti Mini-Extruder five laps at room temperature (see Note 8). 3. Suspend 1 g of Sephadex G-50 resin with 15 mL of solution B. 4. Pour 6 mL of the Sephadex G-50 resin suspension in a disposable plastic gravity flow column (e.g., PD-10, Cytiva). 5. Load the filtered IMV solution on the column. 6. Wash the column with 1 mL of solution B. 7. Add 1.5 mL of solution B and collect the eluate (see Note 9). 8. Adjust the concentration of IMV to give an OD600 value of 0.1.

3.5 Transport Assay (500 μL)

The final assay mixture contains 20 mM Tris–HCl pH 7.5, 125 mM KCl, 5 mM MgCl2, 1 mM dithiothreitol (DTT), 5 mM ATP, 4 μM FlgD (substrate protein), 0.25 μM FliJ, and the 1.5 μM FliH2/FliI complex. 1. Thaw the frozen stock of the IMV solution at room temperature. 2. Mix 100 μL of the IMV solution with 375 μL of external solution in a 1.5 mL polyallomer tube (with cap, for micro ultracentrifuge). 3. Add 25 μL of 0.1 M ATP solution (final conc: 5 mM) to initiate the protein export. Mix well by pipetting ten times using a 1000 μL pipette set at 450 μL. 4. Pick 5 μL of the assay solution to measure the amount of FlhA by immunoblotting (see Note 10). 5. Incubate the assay mixture at 37  C for appropriate time. 6. Add 28 μL of proteinase K solution to the assay mixture (at a final concentration of 50 μg/mL) to terminate the protein transport. Mix well by pipetting ten times using a 1000 μL pipette set at 450 μL.

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7. Incubate the assay mixture at 37  C for 30 min to fully degrade non-transported substrate proteins. 8. Spin the assay mixture (186,000  g, 35 min, 4  C) using a fixed angle rotor (e.g., TLA-55, Beckman Coulter) to precipitate IMVs (see Note 11). 9. Discard the supernatant. 10. Add 900 μL of solution B, and suspend the solution by pipetting to wash IMV. 11. Spin the solution (186,000  g, 8 min, 4  C) using a fixed angle rotor (e.g., TLA-55, Beckman Coulter) to precipitate IMVs. 12. Discard the supernatant. 13. Add 45 μL of 1% (v/v) Triton X100 solution on ice. Place the tube in a sonic bath to completely dissolve IMVs (typically for 30 s). 14. Add 6 μL of trichloroacetate (TCA), and stir the mixture by a Vortex mixer. 15. Incubate the mixture for 40 min on ice. 16. Spin the mixture (20,000  g, 10 min, 4  C) using a fixed angle rotor (e.g., AR015-24, TOMY). 17. Discard the supernatant by using a 1000 μL pipette. 18. Add 100 μL of acetone, and place the tube in a sonic bath for 30 s (see Note 12). 19. Spin the mixture (20,000  g, 10 min, 4  C) using a fixed angle rotor (e.g., AR015-24, TOMY). 20. Discard the supernatant by using a 1000 μL pipette. 21. Add 50 μL of SDS-PAGE sample buffer. 22. Heat the solution on a heat block at 103  C for 3 min. 23. The tubes can be stored at necessary. 3.6 Detection of the Transported Proteins

30  C until before SDS-PAGE, if

1. Apply the sample (~3 μL) to SDS-PAGE. 2. Immunoblotting with polyclonal anti-FlgD (export substrate) antibody. 3. Apply the picked sample (see Subheading 3.5, step 4) to SDSPAGE. 4. Immunoblotting with polyclonal anti-FlhA antibody.

In Vitro Protein Transport Assay

4

25

Notes 1. The inside of IMV is filled by solution A. 2. Expression and purification of the FliH2/FliI complex, FliJ, and FlgD were previously described [16]. 3. This Salmonella mutant cells produce the basal bodies but not the hooks because FlgD is essential for hook assembly [19]. Deletion of FliT and overexpression of FlhD/FlhC from the plasmid increase the number of the flagellar basal bodies [20]. The FlhB(N269A) mutation locks the export apparatus in the mode for rod-hook-type protein export [21]. To utilize purified FlgD for evaluation of the export activity, a ΔflgD allele is used. 4. Completely remove the supernatant, because LB is harmful for spheroplast formation. 5. Completely remove the supernatant by a pipette. 6. Fully suspend the spheroplasts by a pipette. Otherwise, the spheroplasts tend to be broken into membrane fragments. 7. Cool the cylinder of the homogenizer with ice beforehand. 8. Before usage, wash the filter by two laps of the Avanti MiniExtruder with 500 μL of solution A. 9. The fraction containing IMV is cloudy (Fig. 2). 10. FlhA is an essential component of fT3SS and is used to evaluate the amount of fT3SS in the assay mixture. 11. You can add inhibitor of proteinase K. 12. For removal of TCA and Triton X100. Their contaminations often give a bad effect on SDS-PAGE.

Fig. 2 Elution from the Sephadex G-50 column. (a) Washing eluate (clear) and (b) eluate containing IMV (cloudy)

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Acknowledgments This work was supported in part by JSPS KAKENHI Grant Numbers 21H02443, 21H00402, and JP15H02386 to K.I., and 18K07108 and 21K07022 to H.T. References 1. Iino T (1974) Assembly of Salmonella flagellin in vitro and in vivo. J Supramol Struct 2:372– 384 2. Zhao Z, Zhao Y, Zhuang XY et al (2018) Frequent pauses in Escherichia coli flagella elongation revealed by single cell real-time fluorescence imaging. Nat Commun 9:1885 3. Macnab RM (2004) Type III flagellar protein export and flagellar assembly. Biochim Biophys Acta 1694:207–217 4. Minamino T, Imada K, Namba K (2008) Mechanisms of type III protein export for bacterial flagellar assembly. Mol BioSyst 4:1105– 1115 5. Terashima H, Kawamoto A, Morimoto YV et al (2017) Structural differences in the bacterial flagellar motor among bacterial species. Biophys Physicobiol 14:191–198 6. Thomas J, Stafford GP, Hughes C (2004) Docking of cytosolic chaperone-substrate complexes at the membrane ATPase during flagellar type III protein export. Proc Natl Acad Sci U S A 101:3945–3950 7. Imada K, Minamino T, Kinoshita M et al (2010) Structural insight into the regulatory mechanisms of interactions of the flagellar type III chaperone FliT with its binding partners. Proc Natl Acad Sci U S A 107:8812– 8817 8. Minamino T, Kinoshita M, Imada K et al (2012) Interaction between FliI ATPase and a flagellar chaperone FliT during bacterial flagellar protein export. Mol Microbiol 83:168–178 9. Minamino T, Namba K (2008) Distinct roles of the FliI ATPase and proton motive force in bacterial flagellar protein export. Nature 451: 485–488 10. Paul K, Erhardt M, Hirano T et al (2008) Energy source of flagellar type III secretion. Nature 451:489–492 11. Minamino T, Morimoto YV, Hara N et al (2011) An energy transduction mechanism used in bacterial flagellar type III protein export. Nat Commun 2:475

12. Minamino T, Morimoto YV, Kinoshita M et al (2014) The bacterial flagellar protein export apparatus processively transports flagellar proteins even with extremely infrequent ATP hydrolysis. Sci Rep 4:7579 13. Minamino T, Morimoto YV, Kinoshita M et al (2021) Membrane voltage-dependent activation mechanism of the bacterial flagellar protein export apparatus. Proc Natl Acad Sci U S A 118:e2026587118 14. Yamane K, Ichihara S, Mizushima S (1987) In vitro translocation of protein across Escherichia coli membrane vesicles requires both the proton motive force and ATP. J Biol Chem 262: 2358–2362 15. Bageshwar UK, Musser SM (2007) Two electrical potential-dependent steps are required for transport by the Escherichia coli Tat machinery. J Cell Biol 179:87–99 16. Terashima H, Kawamoto A, Tatsumi C et al (2018) In vitro reconstitution of functional Type III protein export and insights into flagellar assembly. mBio 9:e00988–e00918 17. Terashima H, Imada K (2018) Novel insight into an energy transduction mechanism of the bacterial flagellar type III protein export. Biophys Physicobiol 15:173–178 18. Terashima H, Tatsumi C, Kawamoto A et al (2020) In vitro autonomous construction of the flagellar axial structure in inverted membrane vesicles. Biomol Ther 10:126 19. Ohnishi K, Ohto Y, Aizawa S et al (1994) FlgD is a scaffolding protein needed for flagellar hook assembly in Salmonella typhimurium. J Bacteriol 176:2272–2281 20. Aldridge C, Poonchareon K, Saini S et al (2010) The interaction dynamics of a negative feedback loop regulates flagellar number in Salmonella enterica serovar Typhimurium. Mol Microbiol 78:1416–1430 21. Fraser GM, Hirano T, Ferris HU et al (2003) Substrate specificity of type III flagellar protein export in Salmonella is controlled by subdomain interactions in FlhB. Mol Microbiol 48: 1043–1057

Chapter 3 Molecular Simulation to Investigate Open–Close Motion of a Flagellar Export Apparatus Protein FlhAC Akio Kitao Abstract Molecular dynamics (MD) simulation and parallel cascade selection molecular dynamics (PaCS-MD) are widely used to investigate large-amplitude motions of proteins. PaCS-MD is an enhanced conformational sampling method consisting of cycles of parallel unbiased MD simulations combined with a selection of MD snapshots as the initial structures for the next cycle. In addition, free energy calculation can be achieved by the combination of PaCS-MD and the Markov state model (MSM). In this chapter, the protocols to investigate the open–close motion of a flagellar export apparatus protein, FlhAC, by MD and the combination of PaCS-MD and MSM are described. Key words Molecular dynamics simulation, Parallel cascade selection molecular dynamics, Markov state model, Open–close motion, Free energy profile

1

Introduction The type III secretion system (T3SS) of the bacterial flagella transports flagellar structural proteins from the cytoplasm to te distal end of the growing flagellar structure using ATP hydrolysis and proton motive force as the energy sources [1]. The T3SS includes a membrane protein, FlhA, as a part of the export gate complex. FlhA comprises a transmembrane domain (FlhATM), a cytoplasmic domain (FlhAC), and a linker between them [2]. FlhAC forms a nonamer ring [3, 4] that plays various roles in substrate export and energy transduction [5]. FlhAC from Salmonella enterica (S. enterica) consists of four subdomains, D1 (residues 362–434, 484–503), D2 (435–483), D3 (504–583), and D4 (584–692), and shows a large cleft between D2 and D4 (Fig. 1) [2]. An open–close domain motion of FlhAC, characterized by the distance between the centers of mass of domains D2 and D4 (d24), has been observed by all-atom molecular dynamics (MD) simulation of the FlhAC monomer within the time scale of tens of nanoseconds

Tohru Minamino et al. (eds.), Bacterial and Archaeal Motility, Methods in Molecular Biology, vol. 2646, https://doi.org/10.1007/978-1-0716-3060-0_3, © The Author(s), under exclusive license to Springer Science+Business Media, LLC, part of Springer Nature 2023

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Fig. 1 The structure of S. enterica FlhAC (residues 347–692) (PDB ID: 3A5I) [2]. Four subdomains, D1 (residues 362–434, 484–503), D2 (435–483), D3 (504–583), and D4 (584–692), are shown in orange, orange red, green, and light blue, respectively. (Image is created by Chimera X [33, 34])

[2]. Consistently, the open and closed conformations of FlhAC have been shown to exist in crystal states [6, 7]. The S. enterica flhA (G368C) mutant is temperature-sensitive and exports flagellar proteins at 27 °C but not at 42 °C [8]. MD simulations of wild-type FlhAC (WT) and its G368C mutant (G368C) for 1.5 μs have indicated that d24 significantly fluctuates in both WT and G368C at 27 °C. In contrast, at 42 °C, d24 of G368C tends to be restricted to the closed conformation in the last 0.5 μs of the 1.5 μs MD, while that of WT fluctuates largely (Fig. 2a–c) [9]. To further investigate the temperature-dependent open–close motion of FlhAC, free energy profiles of both WT and G368C have been calculated as a function of d24 at both 27 and 42 °C (Fig. 2d) [9]. This free energy calculation is achieved by the combination of parallel cascade selection molecular dynamics (PaCS-MD) and the Markov state model (MSM) [10–13]. PaCS-MD is an unbiased enhanced conformational sampling method conducted as cycles of parallel unbiased MD simulations combined with a selection of MD snapshots as the initial structures for the next cycle [14, 15]. In PaCS-MD, the probability of rare event occurrence toward the quantity for the selection is drastically enhanced by the selection and rerandomization of the initial velocities. In general, the quantity for the selection can be decided depending on the motion to be enhanced, and d24 is employed for simulating the open–close motion of FlhAC [9]. The MSM analysis of multiple short MD trajectories which significantly overlap in conformational space enables us to construct the Markov transition matrix and stationary probabilities of the discretized microstates [16, 17]. This combination, namely, PaCS-MD/MSM, enables us to investigate the free

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Fig. 2 Molecular simulation of WT and G368C. (a, b) Representative structures of WT at 27 °C (blue) and 42 °C (cyan) and G368C at 27 °C (green) and 42 °C (magenta) obtained from 1.5 μs MD. Subdomains D1 and D2 are superimposed. (a) Side and (b) bottom views. (c) Center-of-mass distance between D2 and D4 (d24) during 1.5 μs MD simulation. (d) Free energy change as the function of d24 obtained by PaCS-MD/MSM. Error bars represent the standard error estimated from six independent free energy calculations. (Reprinted from Inoue et al. [9], Copyright (2019), with permission from Elsevier)

energy profile of the open–close motion of FlhAC [9]. The obtained results indicate that the free energy profile is shallow in the wide range of the open–close motion, allowing a large domain motion in WT at both 27 and 42 °C and in G368C at 27 °C, whereas only the closed conformation of G368C has a low free energy at 42 °C, suggesting the confinement of FlhAC in the G368C mutant to the closed form at higher temperature reduces the export function. In this chapter, the protocols to investigate the open–close motion of FlhAC by MD and PaCS-MD/MSM are described.

2

Materials

2.1 Initial Structure for the Wild-Type Simulation

1. Decide the initial structure of a target protein for MD simulation. MD simulation of the WT FlhAC monomer is started from the chain A structure of S. enterica FlhAC (residues 347–692), which has been determined by X-ray crystallography (PDB ID: 3A5I) [2].

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2.2 Initial Structure for the G368C Mutant Simulation

1. Conduct modeling of mutant proteins. For the MD of the G368C mutant, Gly368 is substituted with cysteine by modifying the input PDB file.

2.3 Force Field for the Simulated Systems

1. Select force fields for protein, water, and ions. The AMBER ff99SBildn force field [18] is used for proteins (see Note 1). In the case of the AMBER force fields, more recent versions of the force fields, such as ff14SB [19] and ff19SB [20], are available. WT and G368C are initially solvated with SPC/Eb water molecules [21] and 0.174 M KCl [22].

2.4 Periodic Boundary Condition

1. Decide the simulation box size, place the protein in the box center, and fill the box with water molecules and ions (see Note 1). The simulation boxes are constructed with a margin of at least 12 Å from the proteins to the periodic box boundaries. In this case, a cubic box is employed, and the initial box size is 101 × 101 × 101 Å3.

3

Methods

3.1 Molecular Dynamics Simulation for Equilibration

1. Conduct energy minimization before MD simulation (see Note 2).

3.2 Molecular Dynamics Simulation for Production

1. Perform MD simulation for production. As production runs, MD simulation without restraints should be conducted. MD simulations for 1.5 μs for both the wild-type and mutant FlhAC are performed at 27 and 42 °C.

3.3 Analysis of MD Trajectories

1. Store MD trajectories. For each MD, the last 1.0 μs trajectory is selected, and 10,000 snapshots (every 0.1 ns) are used for the following analyses.

2. Perform equilibration MD with positional restraints for the protein main chain atoms with isothermal-isobaric (NPT) condition at target temperatures and 1 atm, so that protein sidechains, water molecules, and ions are well equilibrated around the protein before allowing free movement of the protein. MD simulations of FlhAC for 1 ns with or without the G368C substitution are independently conducted at 27 and 42 °C.

2. Select representative structures by clustering (see Note 3). The obtained four trajectories (WT and G368C at 27 and 42 °C) are merged into one and clustered into 10 clusters based on the Cα atom RMSD-based K-means clustering. The obtained representative structures from highly populated clusters are selected and presented in Fig. 2a, b. 3. Conduct the principal component analysis (PCA) of the MD trajectories. The merged trajectory is analyzed by PCA as

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shown in the literatures (see Note 4) [23, 24]. Only the Cartesian coordinates of Cα atoms are used in this case. The first principal mode, which is the largest-amplitude motion, is confirmed to be the open–close motion of FlhAC. 3.4 PaCS-MD Simulation of Open– Close Movements

1. Select an appropriate quantity for the selection in PaCS-MD to enhance conformational sampling (see Note 5). Use an interdomain distance between domains D2 and D4, d24, as a parameter. The distance between the centers of mass of the two domains is used to calculate d24. 2. Conduct multiple trials of PaCS-MD for WT and G368C at 27 and 42 °C, so as to sample the conformational space sufficiently. For each case, six independent trials of targeted PaCSMD are conducted. The six trials are conducted from the structures selected from the production MD trajectory at 0.9, 1.0, 1.1, 1.2, 1.3, 1.4, and 1.5 μs MD, respectively. In each trial, two PaCS-MD simulations are performed, targeting either open or closed structures by using d24 as the selection quantity and selecting snapshots with a longer (to open) or shorter (to closed) value of d24, respectively. Each cycle of PaCS-MD consists of 10 independent MD simulations for 2 ns. The PaCS-MD cycle is continued until FlhAC reaches either a completely open or closed structure. The total number of cycles required to reach both open and closed structures is between 30 and 65. In most of the cases, the sampling is completed within 40 cycles.

3.5 Calculation of Free Energy Profile for Open–Close Motion

1. Conduct the MSM analysis (see Note 6). For each trial, snapshots from all the trajectories generated by PaCS-MD are used for the MSM analysis. The first 0.3 ns of each trajectory is excluded from the MSM analysis. 2. Discretize the trajectories. Discretization of the trajectories is conducted with Euclidean metric along the distance space with the minimum distance of 0.2 Å. 3. Decide the best lag time by examining the plot of lag time versus implied timescale. The lag time is set to be 0.1 ns except for one case where 0.2 ns is used. 4. Calculate probability distribution in the stationary state from the Markov transition matrix, and obtain free energy profile as a function of d24. In each case, the obtained stationary probability distribution, p(d24), is interpolated by Spline as the function of d24. The free energy profile is calculated from the average of the six probability distributions as –kBT ln .

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Notes 1. The setup of the simulation system is conducted by the tleap module of AMBER16 [25]. 2. All the simulation is conducted with the pmemd.cuda module [26] of AMBER14 and AMBER16 [25]. A more recent version of AMBER is AMBER21 [27]. 3. K-means clustering is conducted by ClusCo (Clustering and Comparison of Protein Models) [28]. Other clustering libraries can be also used for the clustering of MD trajectories. 4. PCA is conducted by an in-house software. The main calculation of PCA is the diagonalization of the covariance matrix, which can be easily realized as the solution of the standard eigenvalue problem [23, 24]. The numerical calculation can be conducted by math libraries. Also, software for MD trajectory analysis can be used to conduct PCA by, for example, the cpptraj module of AMBER [29]. 5. PaCS-MD can be performed by the MD simulation software of your choice because it is a combination of multiple independent MD simulations. Typically calculation of the selection quantities ranking of snapshots the selection of the initial coordinates and restarting MD simulations with re-randomizing initial velocities can be conducted with a simple script. Use the pmemd.cuda module for each MD simulation as already described (see Note 2) 6. The MSM analysis to calculate the free energy profile is conducted by EMMA 1.4 [30, 31]. A more recent version of this software is available as PyEMMA 2 [32].

Acknowledgments This research was supported by MEXT/JSPS KAKENHI (Nos. JP19H03191, JP20H05439, and JP21H05510) to A.K., and by MEXT as a “Program for Promoting Researches on the Supercomputer Fugaku” (Application of Molecular Dynamics Simulation to Precision Medicine Using Big Data Integration System for Drug Discovery, JPMXP1020200201 and Biomolecular Dynamics in a Living Cell, JPMXP1020200101) to A.K. This work used computational resources of the supercomputer TSUBAME provided by Tokyo Institute of Technology, FUGAKU through the HPCI System Research Project (Project IDs: hp210029, hp210172, and hp210177), and those from Research Center for Computational Science, The National Institute of Natural Science, and The Institute for Solid State Physics, The University of Tokyo.

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References 1. Minamino T (2014) Protein export through the bacterial flagellar type III export pathway. Biochim Biophys Acta 1843:1642–1648 2. Saijo-Hamano Y, Imada K, Minamino T et al (2010) Structure of the cytoplasmic domain of FlhA and implication for flagellar type III protein export. Mol Microbiol 76:260–268 3. Abrusci P, Vergara-Irigaray M, Johnson S et al (2013) Architecture of the major component of the type III secretion system export apparatus. Nat Struct Mol Biol 20:99–104 4. Terahara N, Inoue Y, Kodera N et al (2018) Insight into structural remodeling of the FlhA ring responsible for bacterial flagellar type III protein export. Sci Adv 4:eaao7054 5. Minamino T, Morimoto YV, Hara N et al (2011) An energy transduction mechanism used in bacterial flagellar type III protein export. Nat Commun 2:475 6. Moore SA, Jia Y (2010) Structure of the cytoplasmic domain of the flagellar secretion apparatus component FlhA from Helicobacter pylori. J Biol Chem 285:21060–21069 7. Xing Q, Shi K, Portaliou A et al (2018) Structures of chaperone-substrate complexes docked onto the export gate in a type III secretion system. Nat Commun 9:1773 8. Minamino T, Shimada M, Okabe M et al (2010) Role of the C-terminal cytoplasmic domain of FlhA in bacterial flagellar type III protein export. J Bacteriol 192:1929–1936 9. Inoue Y, Ogawa Y, Kinoshita M et al (2019) Structural insights into the substrate specificity switch mechanism of the type III protein export apparatus. Structure 27:965–976 e966 10. Kitao A, Harada R, Nishihara Y et al (2016) Parallel cascade selection molecular dynamics for efficient conformational sampling and free energy calculation of proteins. AIP Conf Proc 1790:020013 11. Tran DP, Takemura K, Kuwata K et al (2018) Protein-ligand dissociation simulated by parallel cascade selection molecular dynamics. J Chem Theory Comput 14:404–417 12. Tran DP, Kitao A (2019) Dissociation process of a MDM2/p53 complex investigated by parallel cascade selection molecular dynamics and the Markov state model. J Phys Chem B 123: 2469–2478 13. Hata H, Nishihara Y, Nishiyama M et al (2020) High pressure inhibits signaling protein binding to the flagellar motor and bacterial chemotaxis through enhanced hydration. Sci Rep 10: 2351

14. Harada R, Kitao A (2013) Parallel cascade selection molecular dynamics (PaCS-MD) to generate conformational transition pathway. J Chem Phys 139:035103 15. Harada R, Kitao A (2015) Nontargeted parallel cascade selection molecular dynamics for enhancing the conformational sampling of proteins. J Chem Theory Comput 11:5493–5502 16. Pan AC, Roux B (2008) Building Markov state models along pathways to determine free energies and rates of transitions. J Chem Phys 129: 064107 17. Prinz JH, Wu H, Sarich M et al (2011) Markov models of molecular kinetics: generation and validation. J Chem Phys 134:174105 18. Hornak V, Abel R, Okur A et al (2006) Comparison of multiple Amber force fields and development of improved protein backbone parameters. Proteins 65:712–725 19. Maier JA, Martinez C, Kasavajhala K et al (2015) ff14SB: improving the accuracy of protein side chain and backbone parameters from ff99SB. J Chem Theory Comput 11:3696– 3713 20. Tian C, Kasavajhala K, Belfon KAA et al (2020) ff19SB: amino-acid-specific protein backbone parameters trained against quantum mechanics energy surfaces in solution. J Chem Theory Comput 16:528–552 21. Takemura K, Kitao A (2012) Water model tuning for improved reproduction of rotational diffusion and NMR spectral density. J Phys Chem B 116:6279–6287 22. Joung IS, Cheatham TE (2008) Determination of alkali and halide monovalent ion parameters for use in explicitly solvated biomolecular simulations. J Phys Chem B 112:9020–9041 23. Kitao A, Hirata F, Go ¯ N (1991) The effects of solvent on the conformation and the collective motions of protein: Normal mode analysis and molecular dynamics simulations of melittin in water and in vacuum. Chem Phys 158:447– 472 24. Kitao A, Go N (1999) Investigating protein dynamics in collective coordinate space. Curr Opin Struct Biol 9:164–169 25. Case DA, Betz RM, Cerutti DS et al (2016) AMBER 2016. University of California, San Francisco 26. Gotz AW, Williamson MJ, Xu D et al (2012) Routine microsecond molecular dynamics simulations with AMBER on GPUs. 1. Generalized born. J Chem Theory Comput 8:1542–1555

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27. Case DA, Aktulga HM, Belfon K et al (2021) AMBER 2021. University of California, San Francisco 28. Jamroz M, Kolinski A (2013) ClusCo: clustering and comparison of protein models. BMC Bioinf 14:62 29. Roe DR, Cheatham TE 3rd (2013) PTRAJ and CPPTRAJ: software for processing and analysis of molecular dynamics trajectory data. J Chem Theory Comput 9:3084–3095 30. Senne M, Trendelkamp-Schroer B, Mey ASJS et al (2012) EMMA: a software package for Markov model building and analysis. J Chem Theory Comput 8:2223–2238

31. Senne M, Trendelkamp-Schroer B, Mey A et al (2013) EMMA 1.4 Markov model algorithms 32. Scherer MK, Trendelkamp-Schroer B, Paul F et al (2015) PyEMMA 2: a software package for estimation, validation, and analysis of Markov models. J Chem Theory Comput 11: 5525–5542 33. Goddard TD, Huang CC, Meng EC et al (2018) UCSF ChimeraX: meeting modern challenges in visualization and analysis. Protein Sci 27:14–25 34. Pettersen EF, Goddard TD, Huang CC et al (2021) UCSF ChimeraX: structure visualization for researchers, educators, and developers. Protein Sci 30:70–82

Chapter 4 Live-Cell Imaging of the Assembly and Ejection Processes of the Bacterial Flagella by Fluorescence Microscopy Xiang-Yu Zhuang, Chao-Kai Tseng, and Chien-Jung Lo Abstract Bacterial flagella are molecular machines used for motility and chemotaxis. The flagellum consists of a thin extracellular helical filament as a propeller, a short hook as a universal joint, and a basal body as a rotary motor. The filament is made up of more than 20,000 flagellin molecules and can grow to several micrometers long but only 20 nanometers thick. The regulation of flagellar assembly and ejection is important for bacterial environmental adaptation. However, due to the technical difficulty to observe these nanostructures in live cells, our understanding of the flagellar growth and loss is limited. In the last three decades, the development of fluorescence microscopy and fluorescence labeling of specific cellular structure has made it possible to perform the real-time observation of bacterial flagellar assembly and ejection processes. Furthermore, flagella are not only critical for bacterial motility but also important antigens stimulating host immune responses. The complete understanding of bacterial flagellar production and ejection is valuable for understanding macromolecular self-assembly, cell adaptation, and pathogen-host interactions. Key words Bacterial flagellum, Self-assembly, Flagellar ejection

1

Introduction The bacterial flagellum consists of a reversible rotary motor, a short proximal hook, and a thin helical filament [1]. Flagella are not only critical for bacterial motility, but flagellins are also important antigens that can stimulate both the innate inflammatory response and the development of adaptive immunity of the bacterial host [2–4]. The flagellum is made up of about 20 different types of proteins in the final structure [5, 6]. The extracellular flagellar filament is made up of more than 20,000 flagellin subunits. The flagellin molecules are secreted by the flagellar type III secretion system (T3SS) into the flagellar central hollow channel and transported to the distal end (Fig. 1a). The flagellar filament assembly is dynamic, and the observation requires real-time imaging in live cells. Electron microscopy can be applied to imaging the flagellar filaments with nanometer resolution but only in dead cells

Tohru Minamino et al. (eds.), Bacterial and Archaeal Motility, Methods in Molecular Biology, vol. 2646, https://doi.org/10.1007/978-1-0716-3060-0_4, © The Author(s), under exclusive license to Springer Science+Business Media, LLC, part of Springer Nature 2023

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Fig. 1 Schematics of the assembly and ejection processes of the bacterial flagella. (a) The flagellum consists of a thin extracellular helical filament as a propeller, a short hook as a universal joint, and a basal body as a rotary motor. An additional sheath extended from the cell outer membrane covers the flagellar filament in V. alginolyticus. The C-ring is the lower part of the rotor, and the Type III secretion system (T3SS) at the bottom of the C-ring secrets flagellin subunits for filament growth. (b) The ejection of the V. alginolyticus flagellum starts from proteolytic digestion on the rod. The C-ring then mobilizes on the cell inner membrane. Finally, the flagellum is ejected, and the LP-ring is sealed as a relic FOMC

[7]. Fluorescence microscopy with high contrast imaging of labeled proteins is ideal for the flagellar assembly observation [8–10]. In order to achieve fluorescence real-time live-cell measurement of flagellar assembly, fluorescence labeling must be fast compared to the flagellar filament growth rate and low toxic to the cell physiology. A subgroup of bacteria has a sheath covered flagellar filament. The sheath is a membrane-like structure contiguous to the outer membrane [10–13]. Taking the advantage of membrane lipidbilayer properties, the sheath can be easily and quickly labeled using lipophilic fluorescent dyes within a millisecond time scale [10, 14, 15] (Fig. 1a). Therefore, the live-cell flagellar assembly process can be observed in real time [10]. Flagella have been thought of as a permanent complex in the cell. But later a flagellar outer membrane complex (FOMC) was found as a flagellar relic in different bacterial species using cryoelectron tomography [16–18]. A common feature of the FOMC is that it contains the L and P rings, the bushing of the bacterial flagellar motor, with a plug inside the P ring, but does not have the hook, flagellar filament, and MS ring. To catch the flagellar ejection events, Zhuang et al. have applied time-lapse imaging and have shown the polar flagellum of Vibrio alginolyticus being ejected from the cell pole [19, 20]. These single-cell experimental results have confirmed that these bacterial cells actively eject their flagella and leave the FOMC as relics (Fig. 1b). Fluorescence microscopy and easy labeling of sheathed flagella are valuable tools to investigate the assembly and ejection processes of bacterial flagella. In this chapter, we provide detailed protocols for sheathed flagella fluorescence labeling, flagellar growth and assembly measurements, and ejection observations.

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Materials All solutions are prepared using deionized pure water and adjusted to pH 7.5. Prepare and store all reagents at room temperature.

2.1

Solutions

1. VC media: 5.0 g/L Bacto-tryptone, 5.0 g/L Yeast-extract, 4.0 g/L K2HPO4, 30.0 g/L NaCl, 2.0 g/L Glucose. 2. VPG media: 10.0 g/L Bacto-tryptone, 4.0 g/L K2HPO4, 30.0 g/L NaCl, 0.5% (v/v) Glycerol. 3. TMN buffer: 50 mM Tris–HCl, 5 mM MgCl2, 5 mM Glucose, 300 mM NaCl. 4. TMK buffer: 50 mM Tris–HCl, 5 mM MgCl2, 5 mM Glucose, 300 mM KCl.

2.2

Channel Slide

1. Clean cover glass (see Note 1). 2. Double-sided tapes: Scotch 3 M double-sided tape 136 or other double-sided tapes. 3. Poly-L-lysine: 0.1% and 0.0001% poly-L-lysine solution.

2.3 Flagellar Filament Labeling

1. FM 4-64 solution (16 μM FM4-64 in manipulating medium or buffer).

2.4 Microscope Configuration

1. Inverted fluorescence microscope with a high NA (Recommend NA > 1.3) 100× objective (e.g., Nikon TiE with a 100× Apochromat Lambda Oil objective). 2. Fluorescence illumination: High intensity LED illumination (e.g., CoolLED pE-4000). 3. Fluorescence cubes: For flagella observation, choose a 530/30 nm bandpass filter as the excitation filter, a dichroic mirror with 550 nm center wavelength, and a 550 nm longpass emission filter. For C-ring (eGFP-FliG) observation experiment, use a 470 nm/20× bandpass filter as the excitation filter, dichroic mirror with 500 nm center wavelength, a 530 nm/ 40× bandpass emission filter. 4. Camera: High-sensitivity and low-noise cameras such as Electron Multiplying CCD (EMCCD) and Scientific CMOS (sCMOS) cameras. 5. Software: Images acquisition software that can integrate the excitation illumination power, camera exposure time, and timelapse tasks.

2.5

Image Analysis

1. MATLAB, IDL, ImageJ, or compatible software are all suitable for image analysis.

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2.6 V. alginolyticus Strains

1. VIO5 (V. alginolyticus lateral flagellum-defective mutant) [10, 19]. 2. NMB90 (V. alginolyticus long-polar flagellum mutant) [21]. 3. NMB344 (egfp-fliG, derived from VIO5) [19].

3

Methods All of the procedures are carried out at room temperature unless otherwise stated.

3.1 Making Tunnel Slide

1. Use a microscope slide (75 × 26 mm, top side), two strips of double-sided tape, and a clean coverslip (24 × 40 mm, bottom side) to make a tunnel slide as the fluidic chamber (Fig. 2). The cross of the microscope slide and the coverslip make it easy to perform perfusion in the inverted microscope.

3.2 Flagella Growth Measurements

1. Incubate V. alginolyticus strain (from -80 °C, glycerol stock) in 2 mL of VC medium at 30 °C for overnight (16 h) in a shaking incubator. 2. Refresh 20 μL of overnight culture in 1 mL of VPG and 1 mL of TMK for 3 h at 30 °C in the shaking incubator.

Fig. 2 Schematic of a tunnel slide. A simple fluidic chamber consists of a standard microscope slide (top side) and a crossed clean coverslip (bottom side) assembled by two strips of a double-sided adhesive tape. Solution exchange can be achieved by adding solution on one edge of crossed glasses. Cells are added to the space between these two glasses and attached to the poly-L-lysine-coated coverslip surface

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3. The cells are harvested by centrifuge with 2000 g for 2 min. Wash twice with TMK buffer. 4. Load 20 μL of 0.1% poly-L-lysine solution into the flow chamber, and then flush out with 200 μL of TMK buffer. 5. Load 20 μL of the cell suspension into the tunnel slide chamber for 2 min for their immobilization. Unattached cells were flush out with 200 μL of imaging medium (VPG: TMK = 1: 1). 6. Flow 50 μL of the imaging medium with 16 μM FM4-64 into the channel, and then seal the chamber with vacuum grease. 7. Place the sample on the fluorescence microscope (see Note 2). 8. Excite FM 4-64 by CoolLED light source (525 nm), and detect fluorescence with a proper fluorescence cube. 9. Acquire FM 4-64 fluorescence time-lapse images by a sCMOS camera under the control of the camera control software (Nikon NIS-elements) (e.g., Sensitivity: Gain 4, Exposure time: 100 ms, Interval time: 150 s). 10. Image analysis. Extract flagellar filament skeleton by a suitable intensity threshold. Then, use a polynomial function to fit the skeleton pixels, and obtain flagellar 2D projection length (L2D) by the polynomial function integration. The real flagellar length can be calculated by the correction function L3D = c L2D where c is the correction factor (Fig. 3a).

Fig. 3 Image analysis of the flagellar lengths and flagellar growth rates. (a) Extract flagellar filament skeleton by a suitable intensity threshold. Then, use a polynomial function to fit the skeleton pixels and obtain flagellar 2D projection length (L2D) by the polynomial function integration. The real flagellar length can be calculated by the correction function L3D = c L2D where c is the correction factor and can be found by comparing the theoretical L3D and L2D with known helical radius, pitch, and axial length. (b) The flagellar filament 2D projection length (L2D) can be found in each image and converted to real 3D length (L3D). The flagellar filament growth rate is the 3D flagellar length increment (ΔL3D) divided by the observation time interval (ΔT )

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11. Correction factor. Bacterial flagellar filament is a helix, and the fluorescence image is the 2D projection of the 3D helix. Therefore, the length correction is required. The correction factor can be calculated by comparing the real length of the r ffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffi ffi 2 2π∙ðradiusÞ filament L 3D = þ 1∙Z to the 2D projection pitch rffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffi h  i2ffi RZ 2π∙ðradiusÞ 2πz dz , where radius, pitch, L 2D = 0 1 þ ∙ cos pitch pitch and the z are the helix radius, pitch length, and axial length, respectively. For VIO5 and NMB90, the correction factors are 1.08 and 1.02, respectively. 12. Calculate the flagellar growth rate, α, by the length increment ΔL3D over the time interval ΔT (Fig. 3b). α= 3.3 Flagellar Ejection Observation

ΔL 3D ΔT

1. Incubate V. alginolyticus strain NB344 (from -80 °C, glycerol stock) in 2 mL of VC medium at 30 °C for overnight (16 h) in a shaking incubator. 2. Refresh 20 μL of the overnight culture in 2 mL of VC for 3.5 h at 30 °C in the shaking incubator. 3. The cells are harvested by centrifuge with 2000 g for 2 min. Wash twice with TMN buffer. Finally, transfer cells into the desired solutions. 4. Take 20 μL of 0.0001% poly-l-lysine solution into the flow chamber, and then flush with 200 μL TMN buffer (see Note 3). 5. Load 20 μL of the cell suspension into the chamber with 16 μM FM4-64 dye. 6. Excite FM 4-64 and eGFP by CoolLED light source (525 and 470 nm), and detect fluorescence with proper fluorescence cubes, respectively. 7. Acquire flagella fluorescence images by an EMCCD camera under the control of the camera control software (Nikon NIS-elements) (e.g., Sensitivity: 300, Exposure time: 250 ms) (Fig. 4) (see Note 4).

Fig. 4 Flagellar ejection. A demonstration of two-color fluorescence images of the flagellar ejection process. The C-ring (FliG cluster) is detached from the flagellar base. Then the flagellum is released from the cell body

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Notes 1. Put coverslips into saturated KOH in ethanol for 10 min. Rinse them with clean water, and dry them in a clean hood. 2. Be aware of photodamage to bacterial cells. Low excitation is critical for successful live-cell imaging measurements. 3. To observe the flagellar ejection process, the flagella adherence on the glass surface should be prevented using low concentration poly-L lysine coating. 4. For the flagellar ejection experiment, because the timing of the C-ring detachment and the flagellum detachment is different, set the time-lapse duration for 30–60 min for the observation.

Acknowledgments The work was supported by the Ministry of Science and Technology of the Republic of China under contract No. MOST-1092628-M-008-001-MY4. References 1. Macnab RM (2003) How bacteria assemble flagella. Annu Rev Microbiol 57:77–100 2. Honko AN, Mizel SB (2005) Effects of flagellin on innate and adaptive immunity. Immunol Res 33:83–101 3. Salazar-Gonzalez RM, McSorley SJ (2005) Salmonella flagellin, a microbial target of the innate and adaptive immune system. Immunol Lett 101:117–122 4. Miao EA, Andersen-Nissen E, Warren SE et al (2007) TLR5 and Ipaf: dual sensors of bacterial flagellin in the innate immune system. Semin Immunopathol 29:275–288 5. Armitage JP, Berry RM (2020) Assembly and dynamics of the bacterial flagellum. Annu Rev Microbiol 74:181–200 6. Nirody JA, Sun Y, Lo C (2017) The biophysicist’s guide to the bacterial flagellar motor. Adv Phys X 2(2):324–343 7. Iino T (1974) Assembly of Salmonella flagellin in vitro and in vivo. J Supramol Struct 2:372– 384 8. Turner L, Stern AS, Berg HC (2012) Growth of flagellar filaments of Escherichia coli is independent of filament length. J Bacteriol 194: 2437

9. Renault TT, Abraham AO, Bergmiller T et al (2017) Bacterial flagella grow through an injection-diffusion mechanism. elife 6:e23136 10. Chen M, Zhao Z, Yang J et al (2017) Lengthdependent flagellar growth of Vibrio alginolyticus revealed by real time fluorescent imaging. elife 6:e22140 11. Glauert AM, Kerridge D, Horne RW (1963) The fine structure and mode of attachment of the sheathed flagellum of Vibrio metchnikovii. J Cell Biol 18:327–336 12. Allen RD, Baumann P (1971) Structure and arrangement of flagella in species of the genus Beneckea and Photobacterium fischeri. J Bacteriol 107:295–302 13. McCarter LL (2001) Polar flagellar motility of the Vibrionaceae. Microbiol Mol Biol Rev 65: 445–462 14. Grossart HP, Steward GF, Martinez J et al (2000) A simple, rapid method for demonstrating bacterial flagella. Appl Environ Microbiol 66:3632 15. Wu Y, Yeh FL, Mao F et al (2009) Biophysical characterization of styryl dye-membrane interactions. Biophys J 97:101–109 16. Ferreira JL, Gao FZ, Rossmann FM et al (2019) γ-proteobacteria eject their polar flagella under nutrient depletion, retaining

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flagellar motor relic structures. PLoS Biol 17: e3000165 17. Kaplan M, Subramanian P, Ghosal D et al (2019) In situ imaging of the bacterial flagellar motor disassembly and assembly processes. EMBO J 38:e100957 18. Zhu S, Schniederberend M, Zhitnitsky D et al (2019) In situ structures of polar and lateral flagella revealed by cryo-electron tomography. J Bacteriol 201:e00117–e00119

19. Zhuang X, Guo S, Li Z et al (2020) Live-cell fluorescence imaging reveals dynamic production and loss of bacterial flagella. Mol Microbiol 114:279–291 20. Zhuang X, Lo C (2020) Construction and loss of bacterial flagellar filaments. Biomol Ther 10: 1528 21. Furuno M, Atsumi T, Yamada T et al (1997) Characterization of polar-flagellar-length mutants in Vibrio alginolyticus. Microbiology 143:1615–1621

Chapter 5 Purification and CryoEM Image Analysis of the Bacterial Flagellar Filament Tomoko Yamaguchi, Tomoko Miyata, Fumiaki Makino, and Keiichi Namba Abstract The bacterial flagellum is a large assembly of about 30 different proteins and is divided into three parts: the filament that acts as a screw propeller, the hook as a universal joint, and the basal body as a rotary motor. In the case of Salmonella, the filament length is 10–15 μm, which is more than ten times longer than the size of the cell. The filament is composed of only one component protein, flagellin, and is made of 11 protofilaments. The filament can form 12 different supercoiled structures as polymorphic forms. Each protofilament can take either the L (left-handed) or R (right-handed) state, and the number ratio of the protofilaments in these two states determines the shape of the supercoil. Some point mutations in flagellin make the filament straight by making all the protofilaments in one of the two states. The straight filaments enable us to use their helical symmetries for structural analysis by electron cryomicroscopy (cryoEM) and single particle image analysis. Here, we describe the methods for the purification of the flagellar filament and cryoEM data collection and image analysis. Key words Bacterial flagellum, Electron cryomicroscopy, Single particle image analysis

1

Introduction Salmonella infection is one of the major causes of disease involving diarrhea in the world. The Salmonella flagellum is a motility organelle responsible for rapid movement of bacterial cells toward more desirable environments and is critically related to the infection. The flagellum mainly consists of three parts: the basal body that works as a rotary motor; the filament that functions as a screw propeller; and the hook, a universal joint connecting the filament to the motor [1]. The flagellar motor converts the electrochemical potential difference of cations across the cell membrane to mechanical work required for high-speed rotation with almost 100% efficiency, and the maximum rotation speed of the Salmonella flagellar motor has been measured to be 300 revolutions per second (rps) [1, 2], which is as fast as that of the Formula One racing car engine. The motor is powerful enough to rotate the filaments ten times longer than the

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cell length and is able to rotate both clockwise (CW) and counterclockwise (CCW) without changing the direction of ion flow. The Salmonella flagellar filament is a cylindrical tube structure composed of one type of protein, flagellin. Flagellin is composed of four domains: D0, D1, D2, and D3. Domains D0 and D1 form the inner tube, and domains D2 and D3 form the outer tube. The outermost domain D3 is also known to act as the epitope recognized by antibodies of host immune systems [3–5]. The tubular structure of the filament is built up of 11 protofilaments and can form 12 different supercoiled forms to function as a helical propeller. This is called polymorphic supercoiling, which is physically quite interesting because only one type of protein is used to build up the filament structure. The mechanism of polymorphic supercoiling has been explained as follows. Each of the 11 protofilaments can take either of the 2 conformational states, called L (left-handed) and R (right-handed), which are distinct in the axial periodicity to form the protofilament in two different lengths and in the lateral interactions to form different helical lattices of left- and righthanded. Various supercoiled forms of the filament can be generated depending on the number ratio of the protofilaments in the L and R states [6], and the supercoiled form can be changed by mechanical force produced by the switching of motor rotation between CCW and CW. When the cell swims forward, all the flagellar motors rotate CCW, and normally left-handed supercoiled filaments form a bundle behind the cell to produce thrust. When the motors switch their rotation to CW, the filaments change to a right-handed supercoil, and the bundle falls apart so that the cell can tumble and change the swimming direction [7]. If all the protofilaments are either the L or R state, the filament becomes straight called the L-type or R-type straight filament. These straight forms of the filament give a great advantage for structural analysis by electron cryomicroscopy (cryoEM) single particle image analysis. The helical structure of the filament can be described by two specific parameters, the axial rise and twist angel of subunit along the 1-start helix, and we can reconstruct the three-dimensional (3D) structure of the filament from a relatively small number of images by amplifying high-resolution structural information by using helical symmetry with these helical parameters. Here, we describe the methods for purification of the flagellar filament and its cryoEM single particle image analysis. These methods can be applied to the structural analysis of other filamentous structures as well.

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Materials Prepare all solutions using ultrapure water (resistance = 18 MΩ at 25 °C) and analytical grade reagents. Prepare all reagents at room temperature. Store all reagents at 4 °C. Particular attention should be paid to the adequate disposal of material contaminated with Salmonella.

2.1 Bacterial Strain and Plasmid 2.2 Cell Culture and Solution

SJW590: Salmonella expressing only FljB R-type straight filament, fljB(A461V) ΔfliC [8]. 1. SJW590, frozen stock with 20% glycerol, kept at -80 °C. 2. Luria-Bertani (LB) medium: 1% (w/v) Bacto tryptone, 0.5% (w/v) Yeast extract, 0.5% (w/v) NaCl. 3. LB agar plate: 1% (w/v) Bacto tryptone, 0.5% (w/v) Yeast extract, 0.5% (w/v) NaCl, 1.5% (w/v) Bacto agar. 4. 10% sucrose buffer: 100 mM Tris–HCl (pH 7.8), 10% (w/v) sucrose. 5. 100 mM EDTA-HCl (pH 7.8). 6. 10 mg/mL lysozyme: 10 mg/mL lysozyme suspended in 10% sucrose buffer. Prepare just before the experiment. 7. 100 mM MgSO4. 8. 10% (v/v) Triton X-100. 9. 100 mM EDTA-NaOH (pH 11.0). 10. Buffer C: 10 mM Tris–HCl (pH 7.8), 5 mM EDTA-NaOH (pH 7.8), 1% Triton X-100. 11. Buffer A: 20 mM Tris–HCl (pH 7.8), 150 mM NaCl, 1 mM MgCl2. 12. 20% (w/w) and 50% (w/w) sucrose solution in Buffer C. 13. 12.5% precast polyacrylamide gel. 14. 2× Sample buffer solution: 125 mM Tris–HCl (pH 6.8), 4% (w/v) SDS, 20% (w/v) Glycerol, 0.002% (w/v) Bromophenol Blue. 15. Running Buffer: 3 g Tris, 14.4 g Glycine, 1 g SDS per liter. 16. 2% PTA: 2% (w/v) phosphotungstic acid.

2.3 Equipment and Electron Microscope

1. 37 °C incubator. 2. 37 °C heated shaker. 3. 37 °C heated orbital shaker. 4. 200 mL Erlenmeyer flask. 5. 5 L Erlenmeyer flask.

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6. Spectrophotometer. 7. Microcentrifuge. 8. Centrifuge with rotors (e.g., Avanti J-E, JLA-10.500 rotor, JA-20 rotor Beckman Coulter, US). 9. Ultracentrifuge with rotors (e.g., Optima XE-90, 50.2Ti rotor, 70.1Ti rotor, SW 32Ti swinging bucket rotor, Beckman Coulter, US). 10. Centrifuge tubes. 11. Ultracentrifuge tubes (e.g., Open-Top Thinwall Ultra-Clear tube, 25 × 89 mm, Beckman Coulter). 12. Gradient fractionator and gradient master. 13. Fraction collector. 14. Electrophoresis supply. 15. 1.5 mL Eppendorf tubes. 16. Tweezer. 17. Ion coater. 18. Thin layer of continuous carbon-coated copper 200 mesh EM grid. 19. Desiccator. 20. Transmission electron microscope (e.g., JEM-1011, JEOL, Tokyo, Japan). 21. CCD camera (e.g., TemCam-F415A-HS-4 4 k × 4 k CCD, TVIPS, Gauting, Germany). 22. Holey carbon grid (e.g., molybdenum 200 mesh R1.2/1.3 grid, Quantifoil Micro Tools GmbH, Germany). 23. Plunge-freezing device for cryo-grid preparation (e.g.,Vitrobot Mark IV, Thermo Fisher Scientific, US). 24. Transmission electron cryomicroscope (e.g., Titan Krios, Thermo Fisher Scientific, US). 25. Direct electron detector camera (e.g., Falcon II 4 k × 4 k CMOS, Thermo Fisher Scientific, US). 2.4 Program for Data Collection and Structural Analysis by Single-Particle Image Analysis

1. EPU software (Thermo Fisher Scientific, US). 2. MotionCor2 [9]. 3. Gctf v1.06 [10]. 4. RELION-3.0-β2 [11]. 5. PHENIX [12]. 6. UCSF Chimera [13]. 7. MODELLER [14]. 8. Coot [15].

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Methods

3.1 Purification of FljB

1. Streak the Salmonella strain expressing FljB R-type straight flagellar filaments from a frozen stock to an LB plate, and incubate the cells overnight in a 37 °C incubator. 2. Inoculate a single-bacterial colony, and grow the cells overnight in 30 mL of LB medium in a 200 mL Erlenmeyer flask at 37 °C with shaking. 3. Transfer a 13 mL of overnight culture to a 1.3 L volume of LB medium in a 5 L Erlenmeyer flask, and grow the cells to OD600 of 1.0 at 37 °C with orbital shaking incubator (90 rpm). 4. Collect the cells by centrifugation (6700 × g, 10 min, 4 °C, JLA-10.500). 5. Resuspend the cell pellet in 60 mL of 10% sucrose buffer. 6. Slowly add 6 mL of 100 mM EDTA-HCl (pH 7.8) to the cell suspension with stirring on ice (see Note 1). 7. Slowly add 6 mL of 10 mg/mL lysozyme to the cell suspension with stirring on ice (see Note 1). 8. Stir the suspension on ice for 1 h at 4 °C. 9. Add 6 mL of 100 mM MgSO4 and 10% (v/v) Triton X-100 to the suspension, and stir it on ice for 1 h at 4 °C. 10. Add 6 mL of 100 mM EDTA-NaOH (pH 11.0) to the suspension with stirring on ice. 11. Remove the debris by centrifugation (15,000 × g, 20 min, 4 ° C), and collect the supernatant. 12. Adjust the pH to 10.9 with 5 N NaOH to solubilize the remanent outer membrane fragments. Remove undissolved membrane fragments by centrifugation (15,000 × g, 20 min, 4 °C). 13. Collect the flagellar filament (67,000 × g, 60 min, 4 °C).

by

ultracentrifugation

14. Resuspend the sample pellet in 2 mL of Buffer C, and remove the debris by centrifugation (7300 × g, 10 min, 4 °C) and collect the supernatant. 15. Prepare a 20–50% linear sucrose density gradient using a gradient master (see Note 2). Load the supernatant onto the sucrose density gradient, and fractionate the samples by ultracentrifugation (68,000 × g, 14.5 h, 4 °C). 16. Collect the fractions, approximately 700 μL per each tube, using a gradient fractionator and a fraction collector (see Note 3).

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Fig. 1 SDS-PAGE of each fraction collected from the sucrose gradient. The FljB flagellar filament was purified by 20–50% (w/w) sucrose gradient. Numbers on the top indicate the fractions collected by the gradient fractionator and fraction collector (top to bottom order of the centrifugation tube)

17. Check the amount of FljB and contaminants in each fraction by SDS-PAGE (see Note 4 and Fig. 1). 18. Collect the fractions containing the FljB filament (Fig. 1), and dilute it by two- to threefold volume of buffer A. In order to concentrate the sample, ultracentrifuge the solution (104,000 × g, 60 min, 4 °C), and collect the filaments as a pellet. 19. Resuspend the filament pellet in a 10–50 μL of buffer A, and store the sample solution at 4 °C. 3.2 Negative Staining and Sample Observation

1. Place the continuous carbon-coated EM grids on a grass slide, and glow-discharge the grids for 24 s. 2. Dilute the sample solution 10- to 100-fold with buffer A. Mix 5 μL of the sample solution with a same volume of 2% PTA on a Pala film. 3. Place the EM grid on the droplet of mixed solution, and wait for 5–10 s. Pick up the EM grid by tweezer, and remove the excess solution on the grid by a filter paper. 4. Place the stained EM grid on a filter paper, and dry it in a desiccator at least for 30 min. 5. Observe and record the negative stained EM images with transmission electron microscope operating at 100 kV and CCD camera (Fig. 2).

3.3 CryoEM Sample Preparation and Data Collection

1. Place the holey carbon grids on a grass slide, and glowdischarge both sides of the grids for 24 s.

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Fig. 2 Negatively stained EM image of the FljB R-type straight flagellar filament

2. Apply 1.5 μL of the sample solution onto each side of the grid. Using Vitrobot Mark IV, remove the extra solution (blotting time for 3–4 s, 100% humidity, 4 °C), and rapidly plunge the grid into liquid ethane at the liquid nitrogen temperature. 3. Insert the grids into Titan Krios, a transmission electron cryomicroscope operated at 300 kV with a cryo specimen stage cooled with liquid nitrogen. 4. Using a minimum dose system, take all movies with a total dose of 72.1 electron/Å2, a total exposure time of 2 s, and the number of frames of 7, so that an electron dose of each frame is 10.3 electron/Å2. Set a defocus range to 0.2–1.9 μm, and a nominal magnification of ×75,000, corresponding to an image pixel size of 1.06 Å. 3.4 Image Processing and Model Building

1. To avoid the effect of radiation damage and induced movement, use the middle five frames from the second to the sixth for image analysis (a total dose of ~50 e-/Å2). Correct the beam-induced movement and drift by subsequently aligning the five movie frames using MotionCor2 [9]. 2. Estimate the parameters of the contrast transfer function (CTF) for each image using Gctf [10]. 3. Pick up the filaments manually from about 2300 cryoEM images. Segment the filament images in 400 pixels square boxes with 90% overlap, and extract about 200,000 segments with binning 4. Classify them into 40 classes by applying helical symmetry (rise and twist) using two-dimensional (2D) classification in RELION [11] (see Note 5 and Fig. 3).

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Fig. 3 Image processing and structural analysis of the FljB filament. After the motion correction and CTF estimation, the segmented filaments were extracted with binning 4, aligned and classified in 2D. The “good” classes (red squares in the lower left panel) were selected and re-extracted without binning. In the 3D classification process, the low-pass filtered FliC filament was used as an initial model, and only good classes (red square in the middle right panel) were selected. After the first round of 3D auto-refinement, postprocess, and CTF refinement, those processes were repeated until the resolution reached the upper limit

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4. Select the “Good” 2D classes and re-extract them without binning by RELION (Fig. 3). 5. Reconstruct the 3D density map by 3D classification in RELION. Prepare the initial model for 3D classification by applying a 30 Å resolution low-pass filter to the cryoEM density map of the FliC R-type straight flagellar filament [4]. Use this map as an initial model, classify the re-extracted segments into six classes for searching the helical symmetry (rise from 4.6 to 4.9 Å; twist from 64° to 66°), and select good 3D classes (Fig. 3). One map with clear details is used as the initial model in the next step. 6. Refine the density map using selected classes by 3D auto-refine in RELION along with the search of helical symmetry (from 4.8 to 4.9 Å; from 65.8° to 66.0°). 7. Correct the B-factor and estimate the resolution of the map by postprocess in RELION. RELION calculates the crosscorrelation of two half maps generated in 3D auto-refinement process and estimates the resolution of the 3D density map by the “gold standard” Fourier Shell Correlation (FSC) at a 0.143 criterion. 8. To improve the resolution, calculate the CTF of each segment image using CTF refinement in RELION. Recalculate the 3D auto-refinement and postprocess using those CTF refined segments. Repeat this process until the resolution reaches the upper limit (Fig. 3). 9. To build an atomic model, sharpen the density map by PHENIX [12], and visualize the 3D density map using UCSF Chimera [13]. 10. Generate a homology model of FljB from the atomic model of FliC [4] (PDB ID: 1UCU) using MODELLER [14]. Fit the atomic model into the 3D density map by UCSF Chimera. 11. To refine the atomic model, a density map of the FljB monomer was extracted from the density map of the entire FljB filament by extracting the densities within the 4 Å distance from the atoms of one subunit by UCSF Chimera. Refine the atomic model using this density map by PHENIX real-space refinement and Coot [15]. 12. One FljB molecule is surrounded by eight adjacent FljB molecules and interacts with each other. In order to take into account of those interactions, fit the refined FljB atomic model to the density map of the FljB filament, and extract the

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densities of nine FljB molecules by Chimera. Refine the atomic models as described above, and select the central FljB atomic model as a final model. Build and visualize the atomic model of the FljB filament by Chimera.

4

Notes 1. This process should be carried on gently. Each reagent must be added slowly around 1 mL/s to prevent the cell aggregation. It also needs to be carried out at around 4 °C, because the cold shock is necessary for solubilizing the outer membrane in the next step. 2. For preparing a linear sucrose gradient, we use a BioComp marker block to designate the half full point in the centrifuge tube. Place the 50% (w/w) sucrose solution to the bottom of an Open-Top Thinwall Ultra-Clear ultracentrifuge tube, and gently add 20% (w/w) sucrose solution on top of the 50% (w/w) sucrose buffer. Carefully close the tube with the BioComp cap. There are two types of caps, long for large sample volume and short for small, and we prefer to use the short cap. A linear gradient is made by BioComp Gradient Master™. Alternatively, it is also possible to a make linear gradient by tilting the tube horizontally and standing it still for at least 2 h at room temperature. After making the linear gradient, store it at 4 °C until use. 3. Set the piston down speed to 0.09 and the fraction collect time to 25 s. 4. Mix the sample fractions with the sample buffer solution containing 5% of 2-mercaptoethanol at the 1:1 ratio. Boil the sample solutions at 95 °C for 3 min, and centrifuge them for a short time to spin down the condensate. Load the sample solutions and protein standard makers to each lane, and carry out the electrophorese at a current of 30 mA/gel until the sample fractions reach the bottom of the gel. Following the electrophoresis, fix and stain the proteins in the gel by Coomassie Brilliant Blue (CBB). 5. It is possible to pick up the segment images automatically. First, prepare the reference 2D images by picking up dozens of filament images manually, extracting hundreds of segmented images from them, and classifying those images into 2D classes. By using “good” 2D averaged images as the reference, pick up segment images by using RELION Autopick.

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Acknowledgments This work has been supported by JSPS KAKENHI Grant Number JP25000013 (to K.N.) and JP18K06155 (to T.M.). This work has also been supported by Platform Project for Supporting Drug Discovery and Life Science Research (BINDS) from AMED under Grant Number JP19am0101117 to K.N., by the Cyclic Innovation for Clinical Empowerment (CiCLE) from AMED under Grant Number JP17pc0101020 to K.N. and by JEOL YOKOGUSHI Research Alliance Laboratories of Osaka University to K.N. References 1. Berg HC (2003) The rotary motor of bacterial flagella. Annu Rev Biochem 72:19–54 2. Nakamura S, Hanaizumi Y, Morimoto YV et al (2020) Direct observation of speed fluctuations of flagellar motor rotation at extremely low load close to zero. Mol Microbiol 113: 755–765 3. Mimori Y, Yamashita I, Murata K et al (1995) The structure of the R-type straight flagellar filament of Salmonella at 9 Å resolution by electron cryomicroscopy. J Mol Biol 249:69– 87 4. Yonekura K, Maki-Yonekura S, Namba K (2003) Complete atomic model of the bacterial flagellar filament by electron cryomicroscopy. Nature 424:643–650 5. Samatey FA, Imada K, Nagashima S et al (2001) Structure of the bacterial flagellar protofilament and implications for a switch for supercoiling. Nature 410:331–337 6. Asakura S (1970) Polymerization of flagellin and polymorphism of flagella. Adv Biophys 1: 99–155 7. Macnab RM, Ornston MK (1977) Normal-tocurly flagellar transitions and their role in bacterial tumbling. Stabilization of an alternative quaternary structure by mechanical force. J Mol Biol 112:1–30

8. Yamaguchi T, Toma S, Terahara N et al (2020) Structural and functional comparison of Salmonella flagellar filaments composed of FljB and FliC. Biomol Ther 10:246 9. Zheng SQ, Palovcak E, Armache J-P et al (2017) MotionCor2: anisotropic correction of beam-induced motion for improved cryoelectron microscopy. Nat Methods 14:331– 332 10. Zhang K (2016) Gctf: real-time CTF determination and correction. J Struct Biol 193:1–12 11. Zivanov J, Nakane T, Forsberg BO et al (2018) New tools for automated high-resolution cryoEM structure determination in RELION-3. elife 7:e42166 12. Afonine PV, Poon BK, Read RJ et al (2018) Real-space refinement in PHENIX for cryoEM and crystallography. Acta Crystallogr D Struct Biol 74:531–544 13. Pettersen EF, Goddard TD, Huang CC et al (2004) UCSF Chimera – a visualization system for exploratory research and analysis. J Comput Chem 25:1605–1612 14. Webb B, Sali A (2016) Comparative protein structure modeling using MODELLER. Curr Protoc Bioinform 54:5.6.1–5.6.37 15. Emsley P, Lohkamp B, Scott WG et al (2010) Features and development of Coot. Acta Crystallogr D Biol Crystallogr 66:486–501

Part II Flagella-Driven Motility of Bacteria

Chapter 6 Site-Specific Isotope Labeling of FliG for Studying Structural Dynamics Using Nuclear Magnetic Resonance Spectroscopy Tatsuro Nishikino and Yohei Miyanoiri Abstract To understand flagella-driven motility of bacteria, it is important to understand the structure and dynamics of the flagellar motor machinery. We have conducted structural dynamics analyses using solution nuclear magnetic resonance (NMR) to elucidate the detailed functions of flagellar motor proteins. Here, we introduce the analysis of the FliG protein, which is a flagellar motor protein, focusing on the preparation method of the original stable isotope-labeled protein. Key words NMR, Stable isotope labeling, SAIL amino acids, TROSY

1 1.1

Introduction Background

The bacterial flagellar motor is composed of about 30 kinds of proteins and regulates their protein-protein interactions in a spatiotemporal manner, allowing for precise control of cell motility. As a result, bacterial cells move toward more favorable environments for their survival [1, 2]. The rotational direction of the flagellar motor is quickly switched between clockwise (CW) and counterclockwise (CCW) by a device called the C-ring complex [3], which consists of FliG, FliM, and FliN proteins. The direction of flagellar rotation is regulated by chemotactic signals in response to changes in the environments. When the chemotaxis signaling protein CheY is phosphorylated via the chemotactic signaling transduction network, phosphorylated CheY (CheY-P) binds to FliM and FliN in the C-ring, inducing a structural change in the C-ring to allow the motor to switch the direction of flagellar motor rotation from CCW to CW [4–6]. The FliG protein, another component of the

Tohru Minamino et al. (eds.), Bacterial and Archaeal Motility, Methods in Molecular Biology, vol. 2646, https://doi.org/10.1007/978-1-0716-3060-0_6, © The Author(s), under exclusive license to Springer Science+Business Media, LLC, part of Springer Nature 2023

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C-ring complex, has three functional domains, FliGN, FliGM, and FliGC, and interacts with FliM via FliGM [7]. In contrast, FliGC interacts with the stator complex of the flagellar motor and is involved in the generation of rotational torque [8–11]. Therefore, it is considered that the changes in relative orientation between FliGM and FliGC play an important role in the conversion mechanism of flagellar motor rotation via the interaction between CheY-P and FliM [12, 13]. To elucidate the detailed function of FliG in the switch of flagellar motor rotation, we have focused on the FliG protein of marine Vibrio and have identified several variants with defective rotational switch [14, 15]. We have also studied structural dynamics of FliG and its mutant variants using solution nuclear magnetic resonance (NMR) spectroscopy and molecular dynamics simulations [16]. Recently, we succeeded in obtaining epochmaking knowledge about the CW/CCW switching mechanism of the FliGM-FliGC fragment via NMR analysis using Ile and Phe residue-selective isotope-labeled FliG proteins. In this chapter, we introduce a detailed protocol for preparing residue- and stereoselective isotope-labeled NMR samples of FliGM-FliGC fragments using a conventional E. coli protein expression system. 1.2 Overview of the Method

Solution NMR spectroscopy is a powerful method for understanding the structure, interaction, and dynamics of biomolecules, such as proteins and nucleic acids, at atomic resolution. We have reported a three-dimensional structural analysis of various proteins and changes in protein dynamics associated with interactions [16– 21]. However, the weakness of NMR is that the molecular weight of the protein to be analyzed is relatively small (ca. 10–20 kDa). As the molecular weight increases, the number of NMR signals increases and the line width of the signals are broadened, making observation and analysis of the NMR spectrum extremely difficult. Although the development of multidimensional NMR techniques has greatly improved this problem, it remains a major bottleneck in the development of NMR methods. Various isotope labeling techniques have been developed to overcome these problems. Stereo array isotope labeling (SAIL) [22] and methyl-specific isotope labeling techniques [23–31] are widely used for the structural analysis of large molecular proteins and protein complexes. The basic strategy of the SAIL method is to selectively label hydrogen, carbon, and nitrogen atoms in proteins with stable isotopes (e.g., 2H, 13C, and 15N) and efficiently obtain only the necessary NMR information. For this purpose, regio- and stereoselective [2H, 13C, 15N] triple-labeled amino acids (SAIL amino acids) have been synthesized for all 20 types of amino acids that make up proteins. Then, a protein composed of only SAIL amino acids (SAIL protein) has been prepared using a cell-free protein synthesis system [22, 32, 33]. Using the SAIL protein prepared this way, signal overlap and line broadening, which had been serious

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obstacles to the NMR analysis of the commonly used [13C, 15N] uniformly labeled proteins, have significantly been reduced. As a result, precise and rapid three-dimensional structure determination of proteins with a high molecular weight over 40 kDa has been achieved [22]. Although the molecular weight limit of NMR has been improved by the development of the SAIL method, it is still difficult to determine the tertiary structure of proteins and protein complexes larger than 50 kDa. In addition, high-resolution and high-throughput structural determination methods for large molecular proteins have been established using X-ray crystallography and cryo-electron microscopy (cryo-EM). Therefore, in the further development of the NMR method, it is important to capture their local structural dynamics that are difficult to obtain via X-ray and cryo-EM analyses. Under these circumstances, selective isotope labeling techniques for residue-selective functional groups (e.g., amide, methyl, and aromatic moieties) have been widely developed. In general, amino acid residues with a methyl group or an aromatic ring (e.g., Ile, Leu, Val, Ala, Thr, Met, Phe, Tyr, Trp, and His) are abundant in proteins. Therefore, NMR signals derived from these residues are good probes for visualizing the conformational changes and intramolecular interactions of large proteins. Several methods have been established for the preparation of proteins, in which the methyl groups of several amino acid residues are specifically isotope-labeled. In these methods, only the methyl groups of the protein are specifically labeled with 13C and 1H using chemically synthesized isotope-labeled amino acids or amino acid precursors, and all other protons are replaced with deuterium. Using such isotope-labeled proteins, highly sensitive and sharpened 1 H-13C signals are obtained even for 100–1000 kDa proteins [34, 35]. In recent years, improvements in SAIL amino acids have made it possible to observe NMR signals derived from aromatic rings and methylene groups of macromolecular proteins exceeding 100 kDa with high sensitivity [36, 37]. In addition, with the establishment of these labeling techniques, relaxation dispersion methods for methyl groups [25] and quantitative analysis of aromatic ring flipping motion [38], which can provide important information for understanding the structural dynamics of large proteins, have been developed. Therefore, the combination of residue- and stereospecific isotope labeling of methyl and aromatic groups is useful for deciphering the correlation between the structure, dynamics, and molecular mechanisms of flagellar motor protein complexes.

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Materials

2.1 E. coli Cultivation Using a Deuterated M9 Medium

1. E. coli BL21 (DE3) cells transformed with a plasmid encoding the FliGM-FliGC fragment (see Notes 1 and 2). 2. Luria-Bertani (LB) plate: Dissolve 15 g of agar, 10 g of tryptone, 5.0 g of yeast extract, and 5.0 g NaCl in purified water, and dilute to 1 L, and autoclave for 15 min at 121  C. Add appropriate antibiotics after cooling it to around 50  C. 3. LB medium: Dissolve 10 g of tryptone, 5.0 g of yeast extract, and 5.0 g of NaCl in purified water; dilute to 1 L; and autoclave for 15 min. at 121  C. Add appropriate antibiotics after cooling it to approximately 50  C. 4. Deuterium oxide (D2O; D, 99 atom % 2H). 5. Sterile filtration devices: 0.22 μm syringe filter and 0.22 μm filter cup. 6. M9 medium: Dissolve 7.0 g of Na2HPO4, 3.0 g of KH2PO4, 1.0 g of NH4Cl, and 0.50 g of NaCl in purified water; dilute to 987.5 mL; and autoclave it for 15 min at 121  C. After cooling to room temperature, add 1.0 mL of MgSO4 (filter-sterilized 1.0 M solution), 1.0 mL of CaCl2 (filter-sterilized 0.10 M solution), 0.50 mL of filter-sterilized 8.0% (w/v) thiamine, 10 mL of filter-sterilized 20% (w/v) D-glucose, and appropriate antibiotics. 7. Vitamin D2O solution (2000): Dissolve 2.0 mg of inositol, 1.0 mg of folic acid, 1.0 mg of choline chloride, 1.0 mg of nicotinamide, 1.0 mg of D-pantothenic acid calcium salt, 1.0 mg of pyridoxal, and 0.10 mg of riboflavin in 5.0 mL of D2O and filter-sterilize. 8. M9 deuterated medium: Dissolve 7.0 g of Na2HPO4, 3.0 g of KH2PO4, 2.0 g of ammonium chloride (15NH4Cl, 99 atom % 15 N), 1.0 g of 2H D-Glucose (1,2,3,4,5,6,6-D7, 97–98 atom % 2 H), 0.50 g of NaCl, 0.24 g of MgSO4, 20 mg of thiamin hydrochloride, 20 mg of biotin, 11 mg of CaCl2, 6.3 mg of MnCl2, 1.6 mg of FeCl3, 0.50 mL of vitamin D2O solution (2000), and appropriate antibiotics in 1 L of D2O, and then sterilize. 9. Isotope-labeled phenylalanine ([α,β1,δ1,δ2-13C4;β2,ε1,ε2,ζ-2H4;α-15N]-phenylalanine [SAIL-Phe; Fig.1a; see Note 3) for observing 13Cβ-1Hβ3 and 13Cδ#-1Hδ# signals of Phe residues (see Note 3). 10. Isotope-labeled α-ketobutyric acid [methyl-13C, 99 atom % 13 C; 3,3-D2, 98 atom % 2H] α-ketobutyric acid sodium salt (Fig. 1b)] for observing 13Cδ1-1Hδ1* methyl of Ile residues (see Note 4).

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Fig. 1 SAIL phenylalanine and amino acid precursor for isoleucine. The isotopelabeling pattern of (a) L-[α,β1,δ1,δ2-13C4;β2,ε1,ε2,ζ-2H4;α-15N]-phenylalanine (SAIL-Phe) and (b) α-ketobutyric [methyl-13C, 99 atom % 13C; 3,3-D2, 98 atom % 2 H] acid

11. IPTG preparation: Dissolve 1 M stock solution in D2O and filter-sterilize. 12. 1 L shaking flask with baffles. 13. Shaking incubator. 14. Spectrophotometer. 15. Centrifugal separator. 16. Centrifuge tube. 2.2 Protein Purification

1. Metal affinity resin (see Note 5) for hexa-Histidine-tag affinity chromatography. 2. Proteinase inhibitor mixture. 3. Polypropylene column. 4. TN buffer preparation: 50 mM Tris–HCl, pH 7.0, 150 mM NaCl. Sterilize the buffer using a 0.22 μm filter cup. 5. Column wash buffer: 30 mM Imidazole in TN buffer. Sterilize using a 0.22 μm filter cup. 6. Elution buffer: 120 mM Imidazole in TN buffer. Sterilize by a 0.22 μm filter cup. 7. Ultrasonic disintegrator. 8. Size-exclusion chromatography column 200 increase 10/300 GL column).

(e.g., Superdex

9. High-performance liquid chromatography (HPLC) system. 10. Centrifugal filter unit. 2.3 NMR Spectroscopy

1. Deuterium oxide (D2O; D, 99 atom % 2H). 2. Sodium 2,2-Dimethyl-2-silapentane-5-sulfonate (DSS). 3. NMR sample tube (see Note 6). 4. An NMR spectrometer equipped with a 1H optimized tripleresonance cryogenic probe.

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Methods

3.1 Phe and Ile Residue-Specific Isotope Labeling

1. Start with 3 mL of LB medium from a freshly transformed colony of E. coli cells, which overexpress the FliGM-FliGC fragment, and grow the cells at 37  C with shaking at 120–180 rpm. 2. Add 30 μL of the overnight (16–20 h) culture from step 1 to 50% (v/v) M9 deuterated medium (1:1 mixture of M9 medium and M9 deuterated medium), and grow the cells at 37  C with shaking at 120–180 rpm (see Note 7). 3. Add 30 μL of the overnight culture from step 2 to a 3 mL volume of M9 deuterated medium, and grow the cells at 37  C with shaking at 120–180 rpm (see Note 7). 4. Add all of the overnight culture from step 3 to 300 mL of M9 deuterated medium including 1.0 mg of SAIL-Phe in a sterilized shaking flask containing baffles. The starting OD660 should not be less than 0.05. To increase the deuteration rate and prevent excessive aeration, the entire shaking flask is covered with vinyl bags (Fig. 2). Then, the bacteria are grown at 37  C with shaking 120–180 rpm. 5. When OD660 reaches 0.3–0.4 (about 1 h before the induction of protein overexpression), add 14 mg of α-ketobutyric acid (methyl-13C, 99 atom % 13C; 3,3-D2, 98 atom % 2H; final concentration: 70 mg/L) and 2.0 mg of SAIL-Phe (final concentration: 15 mg/L) to the growth medium. Incubation is continued for approximately 1 h in a shaking incubator (see Notes 8 and 9). 6. After confirming that OD660 reached 0.4–0.5, induce protein overexpression with 0.5 mM (final concentration) of IPTG. 7. Continue the growth at 15  C for 9 h with shaking at 120 rpm. 8. Harvest the cells by centrifugation at 5000  g, 4  C, 10 min. 9. Freeze the cell pellet at

3.2 Protein Purification (See Note 10)

80  C until purification.

1. Add protease inhibitor cocktail to a frozen cell pellet to inhibit endogenous proteinase activity. 2. Add 40 mL of TN buffer to the frozen cell pellet, and thaw the cells. 3. Sonicate the suspension to lyse the cells. 4. Remove undisrupted cells by centrifugation at 8000  g and 4  C for 10 min. 5. Ultracentrifuge the supernatant obtained from step 4 at 142,000  g and 4  C for 30 min to separate the insoluble and soluble fractions. 6. Collect the supernatant from step 5.

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Fig. 2 A schematic representation of site-specific isotope-labeling method using conventional E. coli protein expression system for NMR studies

7. Add 5 mL of TN buffer to 5 mg of metal affinity resin in a polypropylene column, and elute buffer to add to the equilibrant resin. This procedure should be repeated twice. 8. Add the soluble fraction from step 6 to the column and elute. 9. Add 1 mL of column wash buffer to the column, and elute to remove nonspecific binding proteins. Repeat this step four times. 10. Add 5 mL of elution buffer to the column, and elute bound proteins. Repeat this step four times. The FliGM-FliGC fragment should be eluted. Elution fractions should be stored at 4  C to avoid degradation. 11. Concentrate elution fractions to 0.25–1.0 mL using an ultrafiltration device. The final sample concentration depends on the column.

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12. Ultracentrifuge the concentrated sample at 108,000  g and 4  C for 10 min to separate the insoluble fraction. 13. Connect a size-exclusion chromatography column (e.g., Superdex 200) to a HPLC system, and equilibrate with TN buffer at least one column volume. 14. Inject the supernatant from step 12 to the column, and elute with TN buffer. Collect the peak fractions at λ280, and store them at 4  C. 3.3 NMR Signal Assignment of Ile δ1 Methyl in FliGM-FliGC

1. After dissolving purified FliGM-FliGC in the appropriate solution buffer, concentrate it to >0.1 mM (volume: 200–500 μL). 2. Add D2O (final concentration, 2–5% (v/v)) and DSS (final concentration, ~0.01% (w/v)) to step 1. Replace it with an NMR sample tube (see Note 6). 3. Record the standard 1H-13C 2D heteronuclear multiple quantum correlation (HMQC) experiment to observe the 1H-13C correlation signal of Ile δ1 methyl (13Cδ1-1Hδ1#) (see Note 11). 4. For the sequence-specific signal assignment, use site-directed mutations of each Ile residue (e.g., Ile-to-Leu mutations). Record the HMQC spectrum for each mutant, and compare it to that of the wild type (i.e., reference spectrum) (Fig. 3). 5. To confirm the signal assignment unambiguously, assign the backbone amide 15NH signal of the Ile residues (see Note 12). Then, conduct 3D 13C NOESY-HMQC to obtain intraresidue NOE between the amide and δ1 methyl of Ile residues (see Note 13). 6. If a 3D structure (or model structure) is available, the interresidue NOE obtained from the 13C NOESY-HMQC

Fig. 3 Sequence-specific signal assignment for Ile d1 methyl in FliGM-FliGC. (a) Unambiguous methyl CH signal assignment for Ile δ1 using single amino acid substitution. The 800 MHz 2D 1H-13C HMQC spectrum of [Ile δ1, Phe δ, β3-13C, 1H; U-2H] FliGM-FliGC (black) and its I238L mutant (red) are shown overlaid. (b) By comparing the spectra of the wild-type and I238L mutant FliGM-FliGC, δ1 the methyl signal from I238 has been identified (dotted line circle). Using several mutant samples, 13 Ile d1 methyl signals in FliGM-FliGC have been identified

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experiment (i.e., NOEs between the δ1 methyl group of Ile residues) provides additional information for the unambiguous signal assignment. 3.4 SequenceSpecific Signal Assignment of SAILPhe in FliGM-FliGC

1. Use the same NMR sample in Subheading 3.3. 2. Record 1H-13C 2D TROSY spectrum [39] to determine the correlation of the 1H-13C signal of methylene (13Cβ-1Hβ3) and aromatic (13Cδ-1Hδ#) CH signals of SAIL-Phe in FliGM-FliGC. 3. After site-directed mutations of each Phe residue (e.g., Phe to Tyr or Phe to Leu mutations), record the 2D TROSY experiment for each mutant, and compare them with those of the wild-type samples (Fig. 4).

Fig. 4 Sequence-specific signal assignment for SAIL-Phe residues in FliGM-FliGC. (a) δ aromatic CH region of the 800 MHz 2D 1H-13C TROSY spectrum of [Ile δ1, Phe δ, β3-13C, 1H; U-2H] FliGM-FliGC, with signal assignment. (b) Unambiguous signal assignment for SAIL-Phe using single amino acid substitution and NOESY. The intra-residue NOE is used for the sequence-specific signal assignment of δ the CH of SAIL-Phe residues in FliGM-FliGC. We have observed the NOE between Hβ3, Hδ, and amide HN in the F254, F256, and F350 residues using [Ile δ1, Phe δ, β3-13C, 1H; U-2H] FliGM-FliGC. (c) Phe (blue) and Ile (red) residues, which have been determined sequence-specific signal assignment, are shown in structure model of Vibrio alginolitycus FliGM-FliGC

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4. To confirm the signal assignment of SAIL-Phe unambiguously, a 3D 13C NOESY-TROSY (or 3D 15N NOESY-TROSY) experiment be conducted to obtain intra-residue NOEs among amide 15NH, methylene 13Cβ-1Hβ3, and aromatic 13 Cδ-1Hδ# (Fig. 4) (see Notes 12 and 14). 5. If a 3D structure (or model structure) is available, the interresidue NOE obtained via the 13C NOESY-HMQC (e.g., The NOE between δ1 the methyl group of Ile and SAIL-Phe) gives additional information for the unambiguous signal assignment.

4

Notes 1. The FliGM-FliGC fragment of Vibrio alginolitycus is cloned into a cold shock vector (e.g., pColdI) [40], which contains a hexahistidine tag at the N-terminus for affinity purification, and the resulting plasmid is transformed into the E. coli BL21 (DE3) strain. For protein expression, other vectors (e.g., T7 promotor-based vector, pET) or general E. coli protein expression strains are probably available. It is recommended that the effective concentration of the inducer for protein expression is screened. 2. The incorporation efficiency of isotope-labeled amino acids could be optimized using auxotrophic E. coli strains [41, 42] instead of the conventional one. 3. Several SAIL aromatic amino acids and aromatic amino acid precursors with different isotopic labeling patterns are commercially available (SAIL Technologies Co. Ltd.). 4. It is also possible to use isotope-labeled isoleucine instead of α-ketobutyric acid. All are commercially available with various isotope-labeling patterns. 5. For histidine-affinity purification, the batch method for Ni-NTA agarose or similar products and prepacked metal affinity column of crude or Ni-NTA agarose resin are also available. The concentration of imidazole in column wash and elution buffers depends on products and proteins. 6. Usually, an NMR tube with an outer diameter (O.D.) of 5 mm is used. If the sample volume is small (200 μL or less), NMR tubes with an O.D. of 3–4 mm or slotted tubes can be used (SHIGEMI Co. Ltd.). 7. These procedures are necessary for adapting E. coli strains to the deuterated M9 medium. If the growth rate of E. coli is extremely slow (e.g., taking 24–48 h to reach OD660 ≒ 0.5), even using this method, it is necessary to optimize the medium composition. For example, adding deuterated amino acids mixture (e.g., Algal amino acid mixture) to deuterated M9

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medium (to the final concentration of 0.5–1.0 g/L) will significantly improve the growth rate. 8. If isotope-labeled isoleucine is used instead of keto acid, add 3 mg of it to the medium (final concentration, 15 mg/L) in divided portions, such as 1 mg at the start of the main culture and 2 mg before induction; the incorporation efficiency to target protein should exceed 95 atom % 13C,2H. 9. When using the amino acid auxotrophic E. coli mutants, the amount of isotope-labeled reagent can be significantly reduced. In the case of the AB2826 strains, which is Phe, Tyr, and Trp auxotrophic [41], only 5 mg/L of the labeled-Phe is required. In contrast, in cultures of the AB2826 strains, Phe, Tyr, and Trp should always be added to the M9 culture medium, whereas stable isotope-labeled ones are used only at the start of the main culture. The incorporation efficiency into the target proteins is approximately 100 atom % 13C,2H. The Ile, Leu, and Val auxotrophic E. coli strains can be used in a similar procedure [42]. 10. General protein purification protocols are also available for uniform and/or site-specific isotope-labeled proteins. Histidine-affinity chromatography could replace other purification methods (e.g., ion-exchange and strept-tag affinity chromatography), depending on protein construction. Importantly, it is recommended that the sample undergoes sizeexclusion chromatography for the final step of purification because preparation of the monodispersed sample is key for improving signal sensitivity and separation for NMR. Cleavage of the fusion tag is also effective for measurements and should be performed before size-exclusion chromatography. 11. By using the SOFAST-HMQC method instead of conventional HMQC, the experiment time can be shortened to approximately one-fifth of the original time [43, 44]. This method is useful for measuring unstable proteins with low solubility (e.g., samples that degrade or precipitate during the experiment). 12. For amide signal assignment, standard triple-resonance NMR is used. Alternatively, systematic amino acid selective isotopelabeling methods [45, 46] allow us to obtain an unambiguous sequence-specific amide signal assignment with simple 2D NMR experiments. 13. The magnetization transfer method between amide and methyl residues (i.e., out-and-back method) is also possible for signal assignment of large molecular proteins [47]. 14. The sensitivity of intra-residue NOE signal between Hβ3 and NH for Phe depends on the χ1 angle, which represents the rotation angle around the Cα-Cβ bond. When χ1 is approximately 60, Hβ3 is located relatively far from NH (~ 3.7 A˚).

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Therefore, the intensity of the NOE signal between them weakens and may not be detected. Such problems can be resolved by specifically observing the Hβ2 signal using a different type of SAIL-Phe (e.g., [α,β1,δ1,δ2-13C4;β3,ε1,ε2,available from SAIL ζ-2H4;α-15N]-phenylalanine; Technologies Co. Ltd). This intra-residue NOE analysis allows us to achieve not only a sequence-specific signal assignment of Phe for residues in large molecular proteins but also their sidechain conformation [48].

Acknowledgments This work was supported by MEXT (Ministry of Education, Culture, Sports, Science and Technology of Japan) Grant-in-Aid for Scientific Research on Innovative Areas Grant number 17H05877 to Y.M., Program for leading Graduate Schools of Japan, Science for the Promotion of Science (17J11237 and 20J00329 to T.N.). T.N. work was supported in part by the Integrative Graduate Education and Research program of Nagoya University. This research was partially supported by Platform Project for Supporting Drug Discovery and Life Science Research (Basis for Supporting Innovative Drug Discovery and Life Science Research (BINDS)) from AMED. References 1. Berg H (2003) The rotary motor of bacterial flagella. Annu Rev Biochem 72:19–54 2. Homma M, Nishikino T, Kojima S (2022) Achievements in bacterial flagellar research with focus on Vibrio species. Microbiol Immunol 66:75–95 3. Francis NR, Sosinsky GE, Thomas D et al (1994) Isolation, characterization and structure of bacterial flagellar motors containing the switch complex. J Mol Biol 235:1261– 1270 4. Welch M, Oosawa KS, Aizawa M et al (1993) Phosphorylation-dependent binding of a signal molecule to the flagellar switch of bacteria. Proc Natl Acad Sci U S A 90:8787–8791 5. Bren A, Eisenbach M (1998) The N terminus of the flagellar switch protein, FliM, is the binding domain for the chemotactic response regulator, CheY. J Mol Biol 278:507–514 6. Dyer CM, Vartanian AS, Zhou H et al (2009) A molecular mechanism of bacterial flagellar motor switching. J Mol Biol 388:71–84 7. Lam KH, Lam WWL, Wong JYK et al (2013) Structural basis of FliG-FliM interaction in

Helicobacter pylori. Mol Microbiol 88:798– 812 8. Zhou J, Lloyd SA, Blair DF (1998) Electrostatic interactions between rotor and stator in the bacterial flagellar motor. Proc Natl Acad Sci U S A 95:6436–6441 9. Lloyd SA, Blair DF (1997) Charged residues of the rotor protein FliG essential for torque generation in the flagellar motor of Escherichia coli. J Mol Biol 266:733–744 10. Yakushi T, Yang J, Fukuoka H et al (2006) Roles of charged residues of rotor and stator in flagellar rotation: comparative study using H +-driven and Na+-driven motors in Escherichia coli. J Bacteriol 188:1466–1472 11. Terashima H, Kojima S, Homma M (2021) Site-directed crosslinking identifies the statorrotor interaction surfaces in a hybrid bacterial flagellar motor. J Bacteriol 203. https://doi. org/10.1128/JB.00016-21 12. Carroll BL, Nishikino T, Guo W et al (2020) The flagellar motor of Vibrio alginolyticus undergoes major structural remodeling during rotational switching. eLife 9:e61446. https:// doi.org/10.7554/eLife.61446

Site-Specific Isotope Labeling of FliG for Studying Structural Dynamics. . . 13. Chang Y, Zhang K, Carroll BL et al (2020) Molecular mechanism for rotational switching of the bacterial flagellar motor. Nat Struct Mol Biol 11:1041–1047 14. Nishikino T, Zhu S, Takekawa N et al (2016) Serine suppresses the motor function of a periplasmic PomB mutation in the Vibrio flagella stator. Genes Cells 21:505–516 15. Nishikino T, Hijikata A, Miyanoiri Y et al (2018) Rotational direction of flagellar motor from the conformation of FliG middle domain in marine Vibrio. Sci Rep 8:17793. https:// doi.org/10.1038/s41598-018-35902-6 16. Miyanoiri Y, Hijikata A, Nishino Y et al (2017) Structural and functional analysis of the C-terminal region of FliG, an essential motor component of Vibrio Na+-driven flagella. Structure 25:1540–1548 17. Katahira M, Miyanoiri Y, Enokizono Y et al (2001) Structure of the C-terminal RNA-binding domain of hnRNP D0 (AUF1), its interactions with RNA and DNA, and change in backbone dynamics upon complex formation with DNA. J Mol Biol 311:973–988 18. Miyanoiri Y, Kobayashi H, Imai T et al (2003) Origin of higher affinity to RNA of the N-terminal RNA-binding domain than that of the C-terminal one of a mouse neural protein, Musashi1, as revealed by comparison of their structures, modes of interaction, surface electrostatic potentials, and backbone dynamics. J Biol Chem 278:41309–41315 19. Yu C, Feng W, Wei Z et al (2009) Myosin VI undergoes cargo-mediated dimerization. Cell 138:537–548 20. Ihara M, Hamamoto S, Miyanoiri Y et al (2013) Molecular bases of multimodal regulation of a fungal transient receptor potential (TRP) channel. J Biol Chem 288:15303– 15317 21. Hou X, Sekiyama N, Ohtani Y et al (2021) Conformational space sampled by domain reorientation of linear diubiquitin reflected in its binding mode for target proteins. Chem Phys Chem 22:1505–1517 22. Kainosho M, Torizawa T, Iwashita Y et al (2006) Optimal isotope labelling for NMR protein structure determinations. Nature 440: 52–57 23. Metzler WJ, Wittekind M, Goldfarb V et al (1996) Incorporation of 1H/13C/15N-{Ile, Leu, Val} into a perdeuterated, 15N-labeled protein: potential in structure determination of large proteins by NMR. J Am Chem Soc 118:6800–6801 24. Gardner KH, Kay LE (1997) Production and incorporation of 15N,13C, 2H (1H-δ1

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42. Miyanoiri Y, Ishida Y, Takeda M et al (2016) Highly efficient residue-selective labeling with isotope-labeled Ile, Leu, and Val using a new auxotrophic E. coli strain. J Biomol NMR 65: 109–119 43. Pervushin K, Vo¨geli B, Eletsky A (2002) Longitudinal 1H relaxation optimization in TROSY NMR Spectroscopy. J Am Chem Soc 124:12898–12902 44. Schanda P, Brutscher B (2005) Very fast two-dimensional NMR spectroscopy for realtime investigation of dynamic events in proteins on the time scale of seconds. J Am Chem Soc 127:8014–8015 45. Kainosho M, Tsuji T (1982) Assignment of the three methionyl carbonyl carbon resonances in Streptomyces subtilisin inhibitor by a carbon13 and nitrogen-15 double-labeling technique. A new strategy for structural studies of proteins in solution. Biochemistry 21:6273–6279 46. Kasai T, Koshiba S, Yokoyama J et al (2015) Stable isotope labeling strategy based on coding theory. J Biomol NMR 63:213–221 47. Tugarinov V, Kay LE (2003) Ile, Leu, Val methyl assignments of the 723-residue malate synthase G using a new labeling strategy and novel NMR methods. J Am Chem Soc 125: 13868–13878 48. Kainosho M, Miyanoiri Y, Terauchi T et al (2018) Perspective: next generation isotopeaided methods for protein NMR spectroscopy. J Biomol NMR 71:119–127

Chapter 7 Site-Directed Cross-Linking Between Bacterial Flagellar Motor Proteins In Vivo Hiroyuki Terashima, Michio Homma, and Seiji Kojima Abstract The bacterial flagellum employs a rotary motor embedded on the cell surface. The motor consists of the stator and rotor elements and is driven by ion influx (typically H+ or Na+) through an ion channel of the stator. Ion influx induces conformational changes in the stator, followed by changes in the interactions between the stator and rotor. The driving force to rotate the flagellum is thought to be generated by changing the stator-rotor interactions. In this chapter, we describe two methods for investigating the interactions between the stator and rotor: site-directed in vivo photo-crosslinking and site-directed in vivo cysteine disulfide crosslinking. Key words Bacterial flagellum, Flagellar motor, Rotary motor, Stator, Rotor, Photo-cross-link, Disulfide cross-link, PomA, MotA, FliG

1 1.1

Introduction Background

The bacterial flagellum serves as a motility organelle to swim in liquid environments or move on solid surfaces. Bacteria rotate the flagella like a screw by a rotary motor embedded in the cell surface, resulting in the generation of propulsion to push or pull the cell body [1–4]. The rotary motor consists of a stator and rotor. The multiple stator units surround the rotor and assemble into or dissociate from the rotor depending on the environmental conditions, such as viscosity [5–11]. The stator unit is composed of MotA and MotB for the H+-driven motor in Escherichia coli and Salmonella enterica, and PomA and PomB for the Na+-driven motor in Vibrio species [12–16]. Recent structural studies have shown that the stator unit forms a 5:2 hetero-heptamer composed of a barrel-like pentamer ring of the A subunit that surrounds the dimer axis of the B subunit [17, 18]. The A subunit has four transmembrane (TM) segments, a cytoplasmic region between the TM2 and TM3 segments and a C-terminal cytoplasmic tail.

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The B subunit has a single TM segment in the N-terminal region and a periplasmic peptidoglycan-binding domain in the C-terminal region [19, 20]. The rotor is composed of the MS-ring embedded in the inner membrane, which is constructed by dozens of FliF subunits, and the C-ring constructed by dozens of FliG and FliM subunits and about a hundred of FliN subunits [21, 22]. The A subunit of the stator assembling around the rotor interacts with FliG in the C-ring. The stator conducts coupling ions (typically H+ or Na+) through an ion channel in the stator complex, which is followed by a change in its conformation [23–25]. The conformational change in the stator influences the stator-rotor interaction, and the change in the interaction is thought to generate torque for flagellar motor rotation. Recent studies have proposed a gear rotation model to describe the rotation mechanism of the flagellar motor [17, 18, 26, 27]. In the model, the stator gear engages the rotor gear, and these two gears rotate continuously. Therefore, revealing the nature of the stator-rotor interaction is important for understanding of the rotation mechanism of the flagellar motor. The interaction network between the stator and rotor has been examined mainly through genetic analyses [28–35]. The findings have shown that conserved charged residues in the stator and rotor proteins are important not only for torque generation but also for stator assembly into the motor. However, biochemical studies to identify the residues involved in the interactions between the stator and rotor have not been conducted in the past three decades [36]. Here, we describe two biochemical methods that we have employed to identify such residues: site-directed photo-crosslinking and site-directed cysteine disulfide cross-linking. 1.2 Overview of Methods

Site-directed in vivo photo-cross-linking is an innovative technique for detecting protein-protein interactions (Fig. 1) [37]. p-BenzoylL-phenylalanine ( pBPA) is a phenylalanine derivative possessing an ultraviolet (UV)-reactive benzophenone group. It is not incorporated into proteins of organisms because there is no specific tRNA or aminoacyl-tRNA synthase for pBPA. However, when an amber suppressor tRNA and a mutated tyrosyl-tRNA synthase derived from Methanococcus jannaschii are expressed in the cell, the mutated tyrosyl-tRNA synthase charges the amber suppressor tRNA with pBPA, which is then incorporated into an amber stop codon in the translation process. Therefore, pBPA can be sitespecifically introduced into the protein of interest by introducing an amber mutation at a targeted amino acid residue. Since the amber stop codon is the least frequently used stop codon in E. coli, it does not have a significant effect on the growth of E. coli. When a protein containing pBPA is UV irradiated, the reactive group in pBPA reacts with a nearby C-H bond. After the UV treatment, the cross-linked products can be detected by immunoblotting. If an interaction partner is already known, the photo-

Cross-Linking Experiments

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Fig. 1 Schematic drawing of site-specific in vivo photo-cross-linking. First, an amber stop codon is introduced into a residue in a gene of interest, and its mRNA is transcribed. Plasmid pEVOL-pBpF expresses a mutated amber stop codon suppressor tRNA and its cognate mutated aminoacyl-tRNA synthase. pBPA is charged into the tRNA by the aminoacyl-tRNA synthase. A pBPA-charged tRNA is incorporated into the amber stop codon in translation. Finally, a pBPA-incorporated protein forms a covalent bond with its interaction partner protein when irradiated with UV

cross-linked product can be examined using an antibody against the partner protein to confirm the interaction. In our previous study, we introduced an amber stop codon into the pomA gene on the plasmid pYS3, performed photo-cross-linking experiments, and were able to investigate the interaction between PomA and FliG using an anti-FliG antibody (Fig. 2). Site-directed in vivo cysteine disulfide cross-linking is a classical technique to detect residue-residue interactions. A pair of candidate residues that may interact with each other are substituted with cysteine. If the thiol groups of the cysteine residues are in close proximity to each other, they form a disulfide cross-link under oxidized conditions. Because the interior of the cell is generally in a reduced condition, the cross-link formation is facilitated by adding an appropriate oxidant such as copper (II) phenanthroline. Since extracellular spaces such as the periplasm are often in an oxidizing condition, crosslink formation may occur spontaneously between cysteine residues located at very close positions. Crosslinked products can be detected by immunoblotting. In our study, we introduced cysteine residues into the pomA and fliG genes on the plasmid pTSK170 and performed disulfide cross-linking experiments. We were thus able to detect the cross-linked products between PomA and FliG using anti-PomA or anti-FliG antibodies (Fig. 3). In this chapter, we describe the methods of the two types of cross-linking experiments performed in E. coli.

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Fig. 2 In vivo photo-cross-linking between V. alginolyticus PomA and E. coli FliG. V. alginolyticus PomA and chimeric PotB were expressed from plasmid pYS3. The amber stop codon suppressor tRNA and the aminoacyl-tRNA synthase were expressed from plasmid pEVOL-pBpF in the E. coli ΔmotAB strain RP6894. The upper and lower panels show immunoblot images produced with antiV. alginolyticus PomA and anti-S. typhimurium FliG antibodies, respectively. The cross-linked products were indicated by black arrowheads. (These images were cited and modified from Terashima et al. [26])

2

Materials All solutions were prepared using pure water or Milli-Q water (Millipore). Broths and PBS buffer were autoclaved at 121 °C for 20 min. Antibiotics and inducers were sterilized using a 0.22 μm pore filter device. The others were dissolved in each solvent. A freezer is usually set to -20 or -30 °C. Room temperature is controlled to about 20 °C (between 15 and 25 °C).

2.1 Site-Directed In Vivo Photo-CrossLinking

1. E. coli strain RP6894 harboring pYS3 with an amber mutation in pomA and pEVOL-pBpF (see Note 1). 2. LB broth: 1% (w/v) tryptone, 0.5% (w/v) yeast extract, 0.5% (w/v) NaCl (see Note 2). 3. TG broth: 1% (w/v) Bacto tryptone (BD), 0.5% (w/v) NaCl, 0.5% (w/v) glycerol (see Note 3). 4. 25 mg/mL chloramphenicol in ethanol. Store in a freezer. 5. 100 mg/mL ampicillin in Milli-Q water. Store in a freezer. 6. 20% (w/v) L-Arabinose in Milli-Q water. Store in a freezer. 7. 0.1 M pBPA in 1 M NaOH solution. Prepare at time of use (see Note 4).

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Fig. 3 In vivo disulfide cross-linking between V. alginolyticus PomA and E. coli FliG. V. alginolyticus PomA, chimeric PotB and E. coli FliG were co-expressed from plasmid pTSK170 in the E. coli ΔmotAΔfliG strain DFB245. The upper and lower panels show immunoblot images produced with anti-V. alginolyticus PomA and anti-S. typhimurium FliG antibodies, respectively. The cross-linked products were indicated by black arrowheads. (These images were cited and modified from Terashima et al. [26])

8. PBS buffer: 137 mM NaCl, 2.7 mM KCl, 10 mM Na2HPO4, 1.76 mM KH2PO4. Store at room temperature. 9. UV lamp (see Note 5). 10. Sodium dodecyl sulfate (SDS) loading buffer: 62.5 mM Tris– HCl pH 6.8, 2% (w/v) SDS, 10% (w/v) glycerol, 0.01% (w/v) bromophenol blue, and 5% (v/v) β-mercaptoethanol. Store in a freezer. 11. Centrifuge. 12. Shaker. 13. 1.5 mL microtubes. 14. 96-well clear microplates. 15. Glass test tubes. 16. Ice. 17. Spectrophotometer. 2.2 Site-Directed In Vivo Cysteine Disulfide Cross-Linking

1. E. coli strain DFB245 harboring pTSK170 with cysteine substitutions in both PomA and FliG (see Note 6). 2. LB broth: 1% (w/v) tryptone, 0.5% (w/v) yeast extract, 0.5% (w/v) NaCl (see Note 2).

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3. TG broth: 1% (w/v) Bacto tryptone, 0.5% (w/v) NaCl, 0.5% (w/v) glycerol (see Note 3). 4. 100 mg/mL Ampicillin in Milli-Q water. Store in a freezer. 5. 20% (w/v) L-Arabinose in Milli-Q water. Store in a freezer. 6. 0.5 M Copper (II) sulfate (CuSO4) in Milli-Q water. Store in a freezer. 7. 1 M phenanthroline in Milli-Q water. Store in a freezer. 8. Working solution of copper phenanthroline: 12 μL of 0.5 M CuSO4, 100 μL of 1 M phenanthroline, 68 μL of PBS per 100 μL of a working solution. Prepare at time of use. 9. 0.2 M N-methylmaleimide (NEM) in ethanol. Store in a freezer. 10. Working solution of NEM: 150 μL of 0.2 M NEM, 850 μL of PBS per 1 mL of a working solution. Prepare at time of use. 11. PBS buffer: 137 mM NaCl, 2.7 mM KCl, 10 mM Na2HPO4, 1.76 mM KH2PO4. Store at room temperature. 12. SDS loading buffer without β-mercaptoethanol: 62.5 mM Tris–HCl pH 6.8, 2% (w/v) SDS, 10% (w/v) glycerol, 0.01% (w/v) bromophenol blue. Store in a freezer. 13. Centrifuge. 14. Shaker. 15. 1.5 mL microtubes. 16. 96-well clear microplates. 17. Glass test tubes. 18. Ice. 19. Spectrophotometer. 2.3 Sample Preparation for SDSPAGE and Immunoblotting

1. Heat block incubator to boil protein samples. 2. SDS-PAGE system. 3. SDS-PAGE running buffer: 25 mM Tris, 192 mM glycine, 1% (w/v) SDS. Store at room temperature. 4. Semi-dry western blotting transfer system. 5. PVDF membrane for immunoblotting. 6. Transfer buffer: 7.27 g of Tris, 33.8 g of glycine in 2400 mL Milli-Q water, and 600 mL of methanol. Store at room temperature. 7. Skim milk. 8. TBS-T buffer: 20 mM Tris–HCl pH 8.0, 500 mM NaCl, 0.05% (w/v) Tween-20. 9. Rabbit anti-V. alginolyticus PomA antibody to detect V. alginolyticus PomA protein.

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10. Rabbit anti-Salmonella typhimurium FliG antibody to detect E. coli FliG protein (see Note 7). 11. Horseradish peroxidase (HRP) conjugated anti-Rabbit IgG antibody. 12. Chemiluminescence imager. 13. ECL immunoblotting signal detection kit.

3

Methods

3.1 Site-Directed In Vivo Photo-CrossLinking

1. Inoculate E. coli cells in LB broth containing 100 μg/mL ampicillin and 25 μg/mL chloramphenicol, and shake a glass test tube at 30 °C overnight (see Note 8). 2. Measure an optical density at 600 nm (OD600) of the overnight culture using a spectrophotometer. 3. Add Ampicillin, chloramphenicol, and pPBA at final concentrations of 100 μg/mL, 25 μg/mL, and 1 mM, respectively, in fresh TG broth. 4. Inoculate the overnight culture of E. coli cells into fresh TG broth, adjust an initial OD600 to 0.1, and shake at 30 °C for 2 h (see Note 9). 5. Two hours later, add L-arabinose at a final concentration of 0.02% (w/v) to express the mutated tyrosyl-tRNA synthase, amber suppressor tRNA from the plasmid pEVOL-pBpF, and the PomA/PotB complex with the amber mutation from the plasmid pYS3 (see Note 10). 6. Shake the culture at 30 °C for 4 h. 7. Measure OD600 of the cell culture. 8. Centrifuge 1.5 mL of the cell culture (3400 × g, 5 min, 4 °C), discard the supernatant using a micropipette, and suspend the cell pellet in 1 mL of PBS buffer. 9. Centrifuge the cell suspension (3400 × g, 5 min, 4 °C), discard supernatant using a micropipette, and resuspend the cell pellet in OD600 × 250 μL of PBS buffer. 10. Put 100 μL of the suspension into a well of a 96-well microplate as an UV-irradiated sample. 11. Put 100 μL of the suspension in a fresh 1.5 mL microtube as an UV-unirradiated sample, and then put on ice. 12. Place the 96-well microplate on ice, and place an UV lamp on the 96-well plate (see Note 11). 13. Illuminate for 5 min using the UV lamp.

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14. Collect the cells into a fresh 1.5 mL microtube, and centrifuge the UV-irradiated and UV-unirradiated samples (3400 × g, 5 min, 4 °C). 15. Discard the supernatant using the micropipette, and suspend the cell pellet in 150 μL of SDS loading buffer. 16. Store the tubes in a freezer until use. 3.2 Site-Directed In Vivo Cysteine Disulfide Cross-Linking

1. Inoculate the E. coli cells in LB broth containing 100 μg/mL ampicillin, and shake the glass test tube at 30 °C overnight (see Note 8). 2. Measure OD600 of spectrophotometer.

the

overnight

culture

using

a

3. Add ampicillin and L-arabinose at final concentrations of 100 μg/mL and 0.02% (w/v), respectively, to fresh TG broth. 4. Inoculate the overnight culture of E. coli cells into a fresh TG broth, adjust an initial OD600 to 0.05, and shake it at 30 °C for 5 h. 5. Measure OD600 of the culture, collect (1000/OD600) μL of the cell culture in a new 1.5 mL microtube, and centrifuge the tube (3400 × g, 5 min, 4 °C). 6. Discard the supernatant using a micropipette, and suspend the cell pellet in 240 μL of the PBS buffer. 7. Add 4 μL of a working solution of copper phenanthroline, and mix well by tapping. 8. Leave for 5 min at room temperature. 9. Add 80 μL of a working solution of NEM to stop oxidation by modifying free thiol groups, and mix well by tapping. 10. Leave for 5 min at room temperature. 11. Centrifuge (3400 × g, 5 min, 4 °C), discard the supernatant using the micropipette, and suspend the cell pellet in 200 μL of SDS loading buffer without β-mercaptoethanol. 12. Boil at 95 °C for 3 min, and put on ice until the samples are cool. 13. Store the tubes in a freezer until use. 3.3 SDS-PAGE and Immunoblot

1. Add 5 μL aliquots of the sample in the photo-cross-linking experiment, or 4 μL aliquots of the sample in the cysteine disulfide cross-linking experiment, for detection of PomA or FliG, respectively, in each lane of an SDS-polyacrylamide gel. 2. Resolve proteins by electrophoresis. 3. Transfer proteins to a PVDF membrane using the semidry western blotting transfer system.

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4. After the transfer, wash the membrane in TBS-T with 3% (w/v) skim milk for more than 30 min at room temperature with gentle shaking. 5. Discard the TBS-T-skim milk solution, and replace with TBS-T-1% (w/v) skim milk. 6. Add each antibody (1:10,000 dilution) to the membrane, and shake gently for more than 1 h at room temperature. 7. Discard TBS-T containing antibody, and wash the membrane twice using TBS-T for 10 min at room temperature. 8. Discard TBS-T, and replace with TBS-T containing 1% (w/v) skim milk. 9. Add HRP conjugated anti-rabbit IgG antibody (1:10,000 dilution) to the membrane, and shake gently for more than 1 h at room temperature. 10. Repeat step 7. 11. Detect signals using a chemiluminescence detection kit and a chemiluminescence imager.

4

Notes 1. We used E. coli strain RP6894 (motA and motB double null mutant) harboring the plasmids pBAD24 (for the vector control, pBR322 origin, ampicillin resistance) or pYS3 (pBAD24 derivative plasmid encoding Vibrio alginolyticus pomA and chimeric potB, of which product is composed of the N-terminal region of V. alginolyticus PomB and the C-terminal region of E. coli MotB), and plasmid pEVOL-pBpF (p15A origin, chloramphenicol resistance) encoding the amber suppressor tRNA and the mutated tyrosyl-tRNA synthase derived from Methanococcus jannaschii. The replication of pBAD24 is derived from pBR322, and that of pEVOL-pBpF is derived from pACYC184; therefore, these two plasmids are compatible with E. coli cells. 2. Although we used a premixed LB broth purchased from Nacalai Tesque (Japan), you can use an in-house prepared LB broth. 3. We usually use Bacto tryptone purchased from BD in the TG broth. 4. We used pBPA purchased from Bachem AG (Switzerland). Since pBPA is dissolved in 1 M sodium hydroxide and added to the TG medium at a 1:100 dilution, the TG medium becomes slightly alkaline. However, pH change does not affect the growth of E. coli. We are not sure whether the stock solution can be stored in a freezer, because we did not compare

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the solution prepared at the time of use with the solution stored in the freezer. 5. We used Model B-100AP UV lamp purchased from Analytik Jena US. 6. We used E. coli strain DFB245 (motA and fliG double null mutant) harboring plasmid pBAD24 or pTSK170 with cysteine substitutions in both PomA and FliG (encoding V. alginolyticus pomA, chimeric potB, and E. coli fliG in pBAD24). 7. Anti-S. typhimurium FliG antibody was a kind gift from Dr. Minamino (Osaka University). The antibody was used for the detection of the E. coli FliG protein because it cross-reacts well with E. coli FliG. 8. We usually chose 30 °C for the temperature of cell culture, not 37 °C, when examining the motility of E. coli, S. typhimurium, or V. alginolyticus. 9. The addition of two antibiotics hindered growth of the E. coli more than the addition of a single antibiotic. Furthermore, the cells were cultured in TG broth, which is a poor medium. Therefore, the cell cultures were initiated at an OD600 of 0.1. 10. It would be better to add L-arabinose in the early-log phase, because expression of amber suppressor tRNA and its pair tRNA synthase may affect the growth of E. coli. 11. Because the UV lamp gets hot, the microplate is usually placed on ice during photo-cross-linking. Because the intensity of UV decreases inversely proportional to the square of the distance from the light source, the UV lamp is placed on top of the microplate to irradiate the sample with UV radiation from as close as possible.

Acknowledgments This work was supported in part by JSPS KAKENHI grant numbers 18K07108 and 21K07022 (to H.T.), 18K19293 (to S.K.), and 20H03220 (to M.H.). References 1. Macnab RM (2003) How bacteria assemble flagella. Annu Rev Microbiol 57:77–100 2. Terashima H, Kojima S, Homma M (2008) Flagellar motility in bacteria structure and function of flagellar motor. Int Rev Cell Mol Biol 270:39–85

3. Morimoto YV, Minamino T (2014) Structure and function of the bi-directional bacterial flagellar motor. Biomol Ther 4:217–234 4. Takekawa N, Imada K, Homma M (2020) Structure and energy-conversion mechanism of bacterial Na+-driven flagellar motor. Trends Microbiol 28:719–731

Cross-Linking Experiments 5. Leake MC, Chandler JH, Wadhams GH et al (2006) Stoichiometry and turnover in single, functioning membrane protein complexes. Nature 443:355–358 6. Reid SW, Leake MC, Chandler JH et al (2006) The maximum number of torque-generating units in the flagellar motor of Escherichia coli is at least 11. Proc Natl Acad Sci U S A 103: 8066–8071 7. Lele PP, Hosu BG, Berg HC (2013) Dynamics of mechanosensing in the bacterial flagellar motor. Proc Natl Acad Sci U S A 110:11839– 11844 8. Tipping MJ, Delaleza NJ, Limb R et al (2013) Load-dependent assembly of the bacterial flagellar motor. mBio 4:e00551-13 9. Antani JD, Gupta R, Lee AH et al (2021) Mechanosensitive recruitment of stator units promotes binding of the response regulator CheY-P to the flagellar motor. Nat Commun 12:5442 10. Pourjaberi SNS, Terahara N, Namba K et al (2017) The role of a cytoplasmic loop of MotA in load-dependent assembly and disassembly dynamics of the MotA/B stator complex in the bacterial flagellar motor. Mol Microbiol 106:646–658 11. Terahara N, Noguchi N, Nakamura S et al (2017) Load- and polysaccharide-dependent activation of the Na+-type MotPS stator in the Bacillus subtilis flagellar motor. Sci Rep 7: 46081 12. Dean GD, Macnab RM, Stader J et al (1984) Gene sequence and predicted amino acid sequence of the motA protein, a membraneassociated protein required for flagellar rotation in Escherichia coli. J Bacteriol 159:991– 999 13. Stader J, Matsumura P, Vacante D et al (1986) Nucleotide sequence of the Escherichia coli MotB gene and site-limited incorporation of its product into the cytoplasmic membrane. J Bacteriol 166:244–252 14. Kojima S, Blair DF (2004) Solubilization and purification of the MotA/MotB complex of Escherichia coli. Biochemistry 43:26–34 15. Asai Y, Kojima S, Kato H et al (1997) Putative channel components for the fast-rotating sodium-driven flagellar motor of a marine bacterium. J Bacteriol 179:5104–5110 16. Sato K, Homma M (2000) Multimeric structure of PomA, the Na+-driven polar flagellar motor component of Vibrio alginolyticus. J Biol Chem 275:20223–20228 17. Santiveri M, Roa-Eguiara A, Ku¨hne C et al (2020) Structure and function of stator units

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of the bacterial flagellar motor. Cell 183:244– 257 18. Deme JC, Johnson S, Vickery O et al (2020) Structures of the stator complex that drives rotation of the bacterial flagellum. Nat Microbiol 5:1553–1564 19. De Mot R, Vanderleyden J (1994) The C-terminal sequence conservation between OmpA-related outer membrane proteins and MotB suggests a common function in both gram- positive and gram-negative bacteria, possibly in the interaction of these domains with peptidoglycan. Mol Microbiol 12:333– 334 20. Kojima S, Imada K, Sakuma M et al (2009) Stator assembly and activation mechanism of the flagellar motor by the periplasmic region of MotB. Mol Microbiol 73:710–718 21. Ueno T, Oosawa K, Aizawa SI (1992) M-ring, S-ring and proximal rod of the flagellar basal body of Salmonella-typhimurium are composed of subunits of a single protein, FliF. J Mol Biol 227:672–677 22. Francis NR, Sosinsky GE, Thomas D et al (1994) Isolation, characterization and structure of bacterial flagellar motors containing the switch complex. J Mol Biol 235:1261– 1270 23. Blair DF, Berg HC (1990) The MotA protein of E. coli is a proton-conducting component of the flagellar motor. Cell 60:439–449 24. Sato K, Homma M (2000) Functional reconstitution of the Na+-driven polar flagellar motor component of Vibrio alginolyticus. J Biol Chem 275:5718–5722 25. Kojima S, Blair DF (2001) Conformational change in the stator of the bacterial flagellar motor. Biochemistry 40:13041–13050 26. Terashima H, Kojima S, Homma M (2021) Site-directed crosslinking identifies the statorrotor interaction surfaces in a hybrid bacterial flagellar motor. J Bacteriol 9:e00016–e00021 27. Carroll BL, Nishikino T, Guo W et al (2020) The flagellar motor of Vibrio alginolyticus undergoes major structural remodeling during rotational switching. elife 9:e61446 28. Zhou JD, Lloyd SA, Blair DF (1998) Electrostatic interactions between rotor and stator in the bacterial flagellar motor. Proc Natl Acad Sci U S A 95:6436–6441 29. Lloyd SA, Blair DF (1997) Charged residues of the rotor protein FliG essential for torque generation in the flagellar motor of Escherichia coli. J Mol Biol 266:733–744 30. Zhou JD, Blair DF (1997) Residues of the cytoplasmic domain of MotA essential for

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torque generation in the bacterial flagellar motor. J Mol Biol 273:428–439 31. Morimoto YV, Nakamura S, Kami-ike N et al (2010) Charged residues in the cytoplasmic loop of MotA are required for stator assembly into the bacterial flagellar motor. Mol Microbiol 78:1117–1129 32. Yorimitsu T, Sowa Y, Ishijima A et al (2002) The systematic substitutions around the conserved charged residues of the cytoplasmic loop of Na+-driven flagellar motor component PomA. J Mol Biol 320:403–413 33. Yorimitsu T, Mimaki A, Yakushi T et al (2003) The conserved charged residues of the C-terminal region of FliG, a rotor component of Na+-driven flagellar motor. J Mol Biol 334: 567–583

34. Yakushi T, Yang J, Fukuoka H et al (2006) Roles of charged residues of rotor and stator in flagellar rotation: comparative study using H+-driven and Na+-driven motors in Escherichia coli. J Bacteriol 188:1466–1472 35. Takekawa N, Kojima S, Homma M (2014) Contribution of many charged residues at the stator-rotor interface of the Na+-driven flagellar motor to torque generation in Vibrio alginolyticus. J Bacteriol 196:1377–1385 36. Tang H, Braun TF, Blair DF (1996) Motility protein complexes in the bacterial flagellar motor. J Mol Biol 261:209–221 37. Chin JW, Martin AB, King DS et al (2002) Addition of a photocrosslinking amino acid to the genetic code of Escherichia coli. Proc Natl Acad Sci U S A 99:11020–11024

Chapter 8 Measurements of the Ion Channel Activity of the Transmembrane Stator Complex in the Bacterial Flagellar Motor Yusuke V. Morimoto and Tohru Minamino Abstract The bacterial flagellum is driven by a rotational motor located at the base of the flagellum. The stator unit complex conducts cations such as protons (H+) and sodium ions (Na+) along the electrochemical potential across the cytoplasmic membrane and interacts with the rotor to generate the rotational force. Escherichia coli and Salmonella have the H+-type stator complex, which serves as a transmembrane H+ channel that couples H+ flow through an ion channel to torque generation whereas Vibrio and some Bacillus species have the Na+-type stator complex. In this chapter, we describe how to measure the ion conductivity of the transmembrane stator complex over-expressed in E. coli cells using fluorescent indicators. Intensity measurements of fluorescent indicators using either a fluorescence spectrophotometer or microscope allow quantitative detection of changes in the intracellular ion concentrations due to the ion channel activity of the transmembrane protein complex. Key words Ion conductivity, Ion channel, Fluorescent protein, Spectroscopy, Fluorescence microscopy, Bacterial flagellar motor

1

Introduction The bacterial flagellum is a motor unit that is employed by many motile bacteria. In the Salmonella bacterial flagellum, the basal body acts as a rotary motor embedded within the membrane at the base of the flagellum, and its rotation is transmitted to the filament, which acts as a helical propeller, via the hook that acts as a universal joint. The motor rotation is powered by proton (H+) motive force (PMF) across the cytoplasmic membrane, and the torque is generated by the interaction between the stator unit, the MotA/B membrane protein complex, and the rotor, the C-ring [1– 6]. The MotA/B stator complex is composed of five MotA and two MotB transmembrane proteins and acts as a transmembrane H+ channel that couples H+ flow through the channel with the

Tohru Minamino et al. (eds.), Bacterial and Archaeal Motility, Methods in Molecular Biology, vol. 2646, https://doi.org/10.1007/978-1-0716-3060-0_8, © The Author(s), under exclusive license to Springer Science+Business Media, LLC, part of Springer Nature 2023

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Fig. 1 Schematic illustration of the active MotA/B stator complex. A highly conserved aspartic acid residue (D33 in Salmonella typhimurium, D24 in Campylobacter jejuni) critical for H+ influx is located in the transmembrane helix of MotB (highlighted in green). A plug segment of MotB (plug) suppresses H+ leakage through the MotA/B complex. The ribbon diagrams show the crystal structure of the PGB domain of S. typhimurium MotB(L119P) dimer (blue, PDB ID: 5Y3Z) and the cryo-EM structure of C. jejuni MotA/B(Δ41–60) (MotB-TM and MotA pentamer are shown in blue and red, respectively. PDB ID: 6YKP) [7, 8]. The size is compared to the cryo-EM structure of the hook-basal body (gray, EMDB ID: EMD-30409) [9]. PG, peptidoglycan layer; IM, inner membrane

generation of torque (Fig. 1) [7, 10, 11]. On the other hand, the PomA/B complex of marine Vibrio spp. and the MotP/S complex of alkaliphilic Bacillus spp. function as the stator of sodium (Na+)driven flagellar motor with high homology to the MotA/B complex [12–14]. The MotA/B complex of the alkaliphilic Bacillus clausii strain has been reported to utilize either H+ or Na+ as the coupling ion depending on the external pH [14]. Interactions between the stator and the rotor are also required for the assembly of the stator unit into the motor [15, 16]. At least 11 MotA/B stator units can be incorporated around a rotor, and the stator complex shows dynamic turnover during motor rotation [17, 18]. The number of incorporated stator units varies with

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changes in the environment, such as ion concentration and external load, and is also correlated with chemotactic responses [19– 21]. The cytoplasmic loop between the transmembrane (TM) helices 2 and 3 of MotA (MotAC) contains highly conserved charged residues, and the electrostatic interactions between MotAC and the rotor protein FliG are critical for flagellar motor rotation [22, 23]. The TM helix of MotB (MotB-TM), together with the TM3 and TM4 helices of MotA, forms a H+ channel. In Salmonella MotB, the highly conserved Asp-33 residue in the MotB-TM is the most critical residue responsible for H+ influx through the MotA/B complex [24–26]. The MotA/B(D33N) complex do not have the H+ channel activity, resulting in a motility-deficient phenotype. The N-terminal tail of MotB is located within the cytoplasm, and the sizeable C-terminal domain containing a well-conserved peptidoglycan binding (PGB) motif is located in the periplasm [27– 29]. The flexible linker connecting MotB-TM and MotB-PGB contains a plug segment that binds to the H+ channel and suppresses the H+ channel activity when the stator is not incorporated into the motor [30, 31]. The interaction between MotA and FliG is hypothesized to cause a conformational change in the N-terminal portion of MotB-PGB from a compact structure to an extended structure, allowing MotB-PGB to reach and bind to the PG layer (Fig. 1) [8, 32]. A fusion of green fluorescence protein (GFP) to the N-terminus of MotB partially facilitates the H+ channel activity of the stator complex, suggesting that the N-terminal tail of MotB is involved in the stator activation mechanism [33]. Overexpression of a plug-deletion mutant variant of MotB with MotA in E. coli cells results in the formation of the MotA/B (Δ52–71) complex, causes a marked decrease in intracellular pH, and arrests the cell growth [30, 31]. In contrast, overexpression of MotA/B(Δ52–71) with the MotB(D33N) substitution does not change the intracellular pH at all. Although not as much as the plug deletion mutant, overexpression of the wild-type MotA/B stator complex increases the intracellular H+ concentration significantly and so reduces the growth rate of E. coli cells [31, 33]. These indicate that the H+ channel activity of the stator complex can be evaluated by monitoring the change in the intracellular ion concentration of E. coli cells overexpressing the MotA/B stator complex. Some fluorescent proteins change their fluorescence intensity depending on pH, and such pH-sensitive fluorescent proteins can be used as intracellular pH indicators to measure intracellular pH in living cells [34–37]. A GFP derivative, pHluorin, with peak excitation wavelength at 395 and 475 nm and emission at 508 nm, is a remarkable ratiometric fluorescent probe to measure the cytoplasmic pH in living cells [34, 35]. The M153R substitution in pHluorin significantly stabilizes its fusion products while retaining the pH dependence of the fluorescence intensity [35, 38]. A Na+sensitive fluorescent dye, CoroNa Green, is useful for the

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measurement of cytoplasmic Na+ concentrations in E. coli cells and is suitable for evaluating the Na+ channel activity of the Na+-type stator complex [39–41]. In this chapter, we provide detailed protocols to detect the ion channel activity of the stator complexes using fluorescent ion indicators.

2

Materials Prepare all solutions using ultrapure water and analytical grade reagents.

2.1 Bacteria Strain and Plasmids

1. E. coli BL21(DE3) cell (Novagen). 2. pBAD24 (arabinose inducible expression vector) (AmpR) [42]. 3. pACTrc (IPTG inducible expression vector) (CmR) [43]. 4. pYC20: pBAD24/S. typhimurium MotA MotB [31]. 5. pYC109: pBAD24/S. (Δ52–71) [31].

typhimurium

MotA

MotB

6. pYC112: pBAD24/S. typhimurium MotA MotB(Δ52–71, D33N) [44]. 7. pYVM132: pACTrc/pHluorin(M153R) (CmR) (This study). 8. pBAD-PomΔplug: pBAD24/V. alginolyticus PomA PomB (Δ41–120) [40]. 2.2

Culture Media

1. L-broth (LB): 10 g Bacto tryptone, 5 g Yeast extract, 5 g NaCl per liter. 2. T-broth: 1% (w/v) Bacto tryptone, 10 mM potassium phosphate, pH 7.0 with or without 100 mM NaCl. 3. Ampicillin sodium. 4. Chloramphenicol. 5. L-arabinose. 6. Isopropyl-β-D(-)-thiogalactopyranoside (IPTG). 7. Shaking incubator (30
C, at 200 rpm). 8. Test tubes. 9. 1.5 mL microtubes. 10. Single channel pipettes (1000, 100 μL). 11. Microcentrifuge (able to hold 1.5 mL tube, spin at 6000  g).

2.3 Fluorescence Spectrophotometry

1. Fluorescence spectrophotometer. 2. Cuvette with four polished windows. 3. Motility buffer: 10 mM potassium phosphate pH 7.0, 0.1 mM EDTA, 10 mM L-sodium lactate.

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4. 1.5 mL microtubes. 5. Single channel pipettes (1000, 100 μL). 6. pH calibration buffer: Motility buffer adjusted to different pH values: 5.5, 6.0, 6.5, 7.0, 7.5, 8.0, or 8.5 with HCl or KOH. 7. Purified His-pHluorin(M153R) protein in PBS (see Note 1). 8. Prism (GraphPad). 9. Microsoft Excel (Microsoft). 2.4 Fluorescence Microscopy

1. Inverted fluorescence microscope. 2. sCMOS camera. 3. 130 W mercury light source system. 4. Fluorescence mirror unit (for GFP or B-excitation). 5. Vibration isolation optical table. 6. 24 mm  32 mm coverslip (thickness: 0.12–0.17 mm). 7. 18 mm  18 mm coverslip (thickness: 0.12–0.17 mm). 8. Double-sided tape. 9. Tweezers. 10. Single channel pipettes (1000, 100 μL). 11. Motility buffer: 10 mM potassium phosphate pH 7.0, 0.1 mM EDTA, 10 mM L-sodium lactate. 12. 1.5 mL microtubes. 13. Aluminum foil. 14. Filter paper. 15. Microcentrifuge (able to hold 1.5 mL tube, spin at 6000  g). 16. CoroNa Green-AM. 17. Tube rotator (able to hold 1.5 mL tubes, rotate at 5 rpm). 18. Ethylenediamine-N,N,N0 ,N0 -tetraacetic acid, dipotassium salt, dihydrate (EDTA 2K). 19. Gramicidin. 20. Carbonyl cyanide 3-chlorophenylhydrazone (CCCP). 21. Image J (National Institutes of Health, https://imagej.nih. gov/ij/) or Fiji [45]. 22. Prism (GraphPad). 23. Microsoft Excel (Microsoft).

3

Methods Carry out all procedures at ca. 23
C unless otherwise specified.

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3.1 Preparation of Bacterial Samples

1. Grow E. coli cells carrying two different plasmids encoding either stator complex or pHluorin(M153R) overnight in 5 mL LB containing 100 μg/mL ampicillin and 30 μg/mL chloramphenicol at 30
C with shaking. 2. Inoculate 50 μL of the overnight culture of E. coli cells into 5 mL of fresh T-broth containing inducers, 0.2% arabinose (w/v), and 0.1 mM IPTG, and incubate at 30
C with shaking for 5 h (see Note 2). 3. Collect the cells from 500 μL of the culture by centrifugation (6000  g, 2 min). 4. Suspend the cell pellet in 1.0 mL of motility buffer. 5. Centrifuge at 6000  g for 2 min. 6. Discard supernatant. 7. Repeat step 4–6 twice. 8. Resuspend the cell pellet in 500 μL of the motility buffer.

3.2 Acquisition of Excitation Spectrum of pHluorin(M153R) Expressing in E. coli Cells

1. Prepare fresh E. coli cells carrying either pACTrc or pYVM132 and either pBAD24, pYC20, pYC109, or pYC112 in motility buffer (pH 7.0) (see Note 3). 2. Dilute 10 μL of each cell suspension into a 1 mL volume of motility buffer (pH 5.5) (see Note 4). 3. Transfer the cell suspension to a cuvette for fluorescence spectrophotometry. 4. Set the cuvette into the cell holder of a fluorescence spectrophotometer. 5. Measure the excitation spectrum of pHluorin(M153R) in E. coli cells using the fluorescence spectrophotometer with wavelength (Excitation/Emission), 350–500/508 nm; slit width (Excitation/Emission), 5.0/5.0 nm; scan speed, 300 nm/min; PMT voltage, 700 V (see Note 5). 6. Measure the excitation spectrum of E. coli cells expressing no pHluorin(M153R) using the fluorescence spectrophotometer with the same setting in step 5. 7. Save the data of the excitation spectrums as an ASCII text file (*.txt) or csv format (*.csv).

3.3

pH Calibration

1. Dilute the purified His-pHluorin(M153R) protein 100-fold (~100 μg/mL) in motility buffer with different pH, 5.5, 6.0, 6.5, 7.0, 7.5, 8.0, or 8.5. 2. Measure the excitation spectrum of purified His-pHluorin (M153R) at each pH with the same setting in measurements of E. coli cells. 3. Open the spectrum data in Microsoft Excel.

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Fig. 2 Calibration curve of His-pHluorin(M153R). The fluorescent intensity ratio R390/470 was calculated at each pH value (pH 5.5–8.5) (green dots). The plots were fitted by a sigmoidal function (green line).

4. Calculate the fluorescence intensity ratio at 390 nm and 470 nm excitation (R390/470) of the purified pHluorin (M153R) at each pH value (see Note 6). 5. Plot the R390/470 values as a function of buffer pH to make the calibration curve over a pH range from 5.5 to 8.5 (Fig. 2). 6. Fit the calibration values by a sigmoidal curve function using the graphing software Prism. 3.4 Calculation of the Cytoplasmic pH

1. Define the fluorescence intensity of pHluorin(M153R) expressed in E. coli cells by subtracting the intensity value of E. coli cells expressing no pHluorin(M153R) as the autofluorescence intensity of E. coli cells. 2. Calculate the fluorescence intensity ratio R390/470 of pHluorin (M153R) expressed in E. coli cells with or without the expression of the MotA/B complex (see Note 7). 3. Estimate the cytoplasmic pH from the R390/470 value of each strain using the calibration curve. 4. Calculate the mean and the standard deviations of cytoplasmic pH values in pH 5.5 buffer from the data of three or more independent experiments (Fig. 3) (see Note 8).

3.5 Acquisition of Fluorescence Images of E. coli Cells Stained with CoroNa Green

1. Prepare fresh E. coli cells carrying plasmids pBAD24 or pBADPomΔplug in motility buffer with or without 100 mM NaCl (pH 7.0) (see Note 9). 2. Resuspend the cells in 100 μL of motility buffer (pH 7.0) containing 40 μM CoroNa Green-AM and 10 mM EDTA (see Note 10). 3. Rotate the tubes covered with aluminum foil at 5 rpm for 60 min using a tube rotator. 4. Centrifuge at 6000  g for 2 min.

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Fig. 3 H+ channel activity of the MotA/B stator complex. The cytoplasmic pH of E. coli cells in pH 5.5 buffer were determined using pHluorin. Vertical bars indicate standard deviations. These data were acquired using pHluorin, not pHluorin(M153R). (Modified from Suzuki et al. [46])

5. Discard supernatant, and suspend the cell pellet in 1000 μL of fresh motility buffer. 6. Centrifuge at 6000  g for 2 min. 7. Repeat steps 5 and 6 two more times. 8. Resuspend the cells in 500 μL of fresh motility buffer. 9. Prepare a tunnel slide by sandwiching double-sided tape between 24  32 mm coverslip (bottom) and 18  18 mm coverslip (top) [41, 47]. 10. Add the cell suspension to the tunnel slide and leave for 5–10 min. 11. Wash out unbound cells by adding 100 μL of fresh motility buffer. 12. Absorb the excess amount of the buffer with a piece of filter paper (see Note 11). 13. Put the prepared cells on an inverted fluorescence microscope. 14. Excite CoroNa Green by a 130 W mercury lump, and detect fluorescence with a fluorescence mirror unit (Excitation BP 460–480; Emission BP 495–540) (see Note 12). 15. Acquire CoroNa Green fluorescence images by a sCMOS camera (e.g., Exposure time: 100 ms). 16. Measure the fluorescence intensity of the cells stained with CoroNa Green in motility buffer with 0 mM, 5 mM, 10 mM, 20 mM, 50 mM, or 100 mM NaCl in the presence of 20 μM gramicidin and 5 μM CCCP to plot a calibration curve.

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1. Open the fluorescent images by an image analysis software, ImageJ or Fiji (see Note 13). 2. Apply a rectangular mask for the fluorescent image of the bacterial cell body of 8  8 pixels to the ROI (region of interest). 3. Define the instrumental background intensity as the mean pixel intensity within the ROI of a nearby cell-less region. 4. Subtract the total background intensity from each pixel value. 5. Measure the intensity of CoroNa Green from more than 100 cells under each condition. 6. Plot the calibration curve from the data acquired at various external Na+ concentrations in the presence of 20 μM gramicidin and 5 μM CCCP (see Note 14). 7. Fit the calibration values by a Hill slope function using the graphing software Prism. 8. Estimate the intracellular Na+ concentration from the fluorescent intensity of CoroNa Green of each cell using the calibration curve. 9. Calculate the mean and the standard errors of intracellular Na+ concentration in the buffer with or without 100 mM NaCl from the data of 100 cells or more (Fig. 4).

4

Notes 1. The His10-pHluorin(M153R) protein is purified by nickelnitrilotriacetic acid (Ni-NTA) affinity chromatography from

Fig. 4 Na+ channel activity of the PomA/B stator complex. The cytoplasmic Na+ concentrations of E. coli cells were determined using CoroNa Green. Vertical bars indicate standard errors. These data were acquired in T-broth, not in motility buffer. (Modified from Minamino et al. [40])

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the soluble fractions of E. coli BL21(DE3) cells overexpressing His10-pHluorin(M153R) [38]. 2. It is preferable to culture using T-broth instead of LB to reduce autofluorescence, which interferes with fluorescence measurements. 3. The MotA/B complex with either the plug deletion or the MotB(D33N) substitution is used as a positive or negative control for the H+ channel activity of the MotA/B complex, respectively. 4. Since the cytoplasmic pH of E. coli cells is maintained at about 7.1–7.3 [31, 38], the H+ channel activity becomes more apparent when the extracellular pH is set to 5.5. 5. Since the ratio of the two excitation peaks of pHluorin (M153R) changes depending on the pH, the excitation spectrum is used for pH measurement. 6. Fluorescence intensity values at wavelengths shifted from the peak by about 5–10 nm are used because they show more stable values. 7. Since an expression leakage occurs from the arabinose promotor albeit very small even in the absence of arabinose, the empty vector is a more obvious negative control. 8. To compare the H+ channel activity of wild-type MotA/B and its mutants, their expression level should be confirmed by immunoblotting and corrected for the expression level if they are different. 9. The Na+ channel activity of the transmembrane Na+ channel can be compared with and without NaCl. 10. To allow E. coli cells to incorporate CoroNa Green, 10 mM EDTA is required. To avoid Na+ contamination, EDTA-2K should be used (do not use EDTA-2Na). 11. Filter papers that do not contain fluorescent components should be used to reduce background intensity. 12. The fluorescent filter set for GFP is suitable for CoroNa Green. 13. Fiji is an image processing package based on ImageJ and includes plug-ins specifically for life sciences. 14. Gramicidin and CCCP equilibrate Na+ concentrations inside and outside E. coli cells.

Acknowledgments We thank Keiichi Namba and Takuo Yasunaga for continuous support and encouragement, Michio Homma for kind gift of a plasmid encoding pBAD-PomΔplug, and Gero Miesenbo¨ck for a

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gift of the pHluorin probe. This research has been supported in part by JSPS KAKENHI Grant Numbers JP21K06099 and JP21H05532 (to Y.V.M.), JP19H03182, JP22H02573, and JP22K19274 (to T.M.), MEXT KAKENHI Grant Number JP20H05532 and JP22H04844 (to T.M.), and JST PRESTO Grant Number JPMJPR204B (to Y.V.M.). References 1. Berg HC (2003) The rotary motor of bacterial flagella. Annu Rev Biochem 72:19–54 2. Kojima S, Blair DF (2004) The bacterial flagellar motor: structure and function of a complex molecular machine. Int Rev Cytol 233:93–134 3. Sowa Y, Berry RM (2008) Bacterial flagellar motor. Q Rev Biophys 41:103–132 4. Morimoto YV, Minamino T (2014) Structure and function of the bi-directional bacterial flagellar motor. Biomolecules 4:217–234 5. Nakamura S, Minamino T (2019) Flagelladriven motility of bacteria. Biomolecules 9:279 6. Morimoto YV, Minamino T (2021) Architecture and assembly of the bacterial flagellar motor complex. Subcell Biochem 96:297–321 7. Santiveri M, Roa-Eguiara A, Kuhne C et al (2020) Structure and function of stator units of the bacterial flagellar motor. Cell 183:244– 257.e16 8. Kojima S, Takao M, Almira G et al (2018) The helix rearrangement in the periplasmic domain of the flagellar stator B subunit activates peptidoglycan binding and ion influx. Structure 26: 590–598.e5 9. Yamaguchi T, Makino F, Miyata T et al (2021) Structure of the molecular bushing of the bacterial flagellar motor. Nat Commun 12:4469 10. Deme JC, Johnson S, Vickery O et al (2020) Structures of the stator complex that drives rotation of the bacterial flagellum. Nat Microbiol 5:1553–1564 11. Hu H, Santiveri M, Wadhwa N et al (2022) Structural basis of torque generation in the bi-directional bacterial flagellar motor. Trends Biochem Sci 47:160–172 12. Fujinami S, Terahara N, Krulwich TA et al (2009) Motility and chemotaxis in alkaliphilic Bacillus species. Future Microbiol 4:1137– 1149 13. Li N, Kojima S, Homma M (2011) Sodiumdriven motor of the polar flagellum in marine bacteria Vibrio. Genes Cells 16:985–999 14. Terahara N, Namba K, Minamino T (2020) Dynamic exchange of two types of stator units in Bacillus subtilis flagellar motor in response to

environmental changes. Comput Struct Biotechnol J 18:2897–2907 15. Morimoto YV, Nakamura S, Kami-ike N et al (2010) Charged residues in the cytoplasmic loop of MotA are required for stator assembly into the bacterial flagellar motor. Mol Microbiol 78:1117–1129 16. Morimoto YV, Nakamura S, Hiraoka KD et al (2013) Distinct roles of highly conserved charged residues at the MotA-FliG interface in bacterial flagellar motor rotation. J Bacteriol 195:474–481 17. Leake MC, Chandler JH, Wadhams GH et al (2006) Stoichiometry and turnover in single, functioning membrane protein complexes. Nature 443:355–358 18. Reid SW, Leake MC, Chandler JH et al (2006) The maximum number of torque-generating units in the flagellar motor of Escherichia coli is at least 11. Proc Natl Acad Sci U S A 103: 8066–8071 19. Minamino T, Terahara N, Kojima S et al (2018) Autonomous control mechanism of stator assembly in the bacterial flagellar motor in response to changes in the environment. Mol Microbiol 109:723–734 20. Antani JD, Gupta R, Lee AH et al (2021) Mechanosensitive recruitment of stator units promotes binding of the response regulator CheY-P to the flagellar motor. Nat Commun 12:5442 21. Naaz F, Agrawal M, Chakraborty S et al (2021) Ligand sensing enhances bacterial flagellar motor output via stator recruitment. eLife 10: e62848 22. Zhou J, Blair DF (1997) Residues of the cytoplasmic domain of MotA essential for torque generation in the bacterial flagellar motor. J Mol Biol 273:428–439 23. Zhou J, Lloyd SA, Blair DF (1998) Electrostatic interactions between rotor and stator in the bacterial flagellar motor. Proc Natl Acad Sci U S A 95:6436–6441 24. Sharp LL, Zhou J, Blair DF (1995) Tryptophan-scanning mutagenesis of MotB, an integral membrane protein essential for

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flagellar rotation in Escherichia coli. Biochemistry 34:9166–9171 25. Zhou J, Sharp LL, Tang HL et al (1998) Function of protonatable residues in the flagellar motor of Escherichia coli: a critical role for Asp 32 of MotB. J Bacteriol 180:2729–2735 26. Che YS, Nakamura S, Kojima S et al (2008) Suppressor analysis of the MotB(D33E) mutation to probe bacterial flagellar motor dynamics coupled with proton translocation. J Bacteriol 190:6660–6667 27. De Mot R, Vanderleyden J (1994) The C-terminal sequence conservation between OmpA-related outer membrane proteins and MotB suggests a common function in both gram-positive and gram-negative bacteria, possibly in the interaction of these domains with peptidoglycan. Mol Microbiol 12:333–334 28. Kojima S, Furukawa Y, Matsunami H et al (2008) Characterization of the periplasmic domain of MotB and implications for its role in the stator assembly of the bacterial flagellar motor. J Bacteriol 190:3314–3322 29. Kojima S, Imada K, Sakuma M et al (2009) Stator assembly and activation mechanism of the flagellar motor by the periplasmic region of MotB. Mol Microbiol 73:710–718 30. Hosking ER, Vogt C, Bakker EP et al (2006) The Escherichia coli MotAB proton channel unplugged. J Mol Biol 364:921–937 31. Morimoto YV, Che Y-S, Minamino T et al (2010) Proton-conductivity assay of plugged and unplugged MotA/B proton channel by cytoplasmic pHluorin expressed in Salmonella. FEBS Lett 584:1268–1272 32. Terahara N, Kodera N, Uchihashi T et al (2017) Na+-induced structural transition of MotPS for stator assembly of the Bacillus flagellar motor. Sci Adv 3:eaao4119 33. Morimoto YV, Namba K, Minamino T (2020) GFP fusion to the N-terminus of MotB affects the proton channel activity of the bacterial flagellar motor in Salmonella. Biomolecules 10: 1255 34. Miesenbo¨ck G, De Angelis DA, Rothman JE (1998) Visualizing secretion and synaptic transmission with pH-sensitive green fluorescent proteins. Nature 394:192–195 35. Morimoto YV, Kojima S, Namba K et al (2011) M153R mutation in a pH-sensitive green fluorescent protein stabilizes its fusion proteins. PLoS One 6:e19598

36. Bencina M (2013) Illumination of the spatial order of intracellular pH by genetically encoded pH-sensitive sensors. Sensors (Basel) 13:16736–16758 37. Liu A, Huang X, He W et al (2021) pHmScarlet is a pH-sensitive red fluorescent protein to monitor exocytosis docking and fusion steps. Nat Commun 12:1413 38. Morimoto YV, Kami-Ike N, Miyata T et al (2016) High-resolution pH imaging of living bacterial cells to detect local pH differences. mBio 7:e01911-16 39. Lo CJ, Leake MC, Berry RM (2006) Fluorescence measurement of intracellular sodium concentration in single Escherichia coli cells. Biophys J 90:357–365 40. Minamino T, Morimoto YV, Hara N et al (2016) The bacterial flagellar type III export gate complex is a dual fuel engine that can use both H+ and Na+ for flagellar protein export. PLoS Pathog 12:e1005495 41. Morimoto YV, Namba K, Minamino T (2017) Bacterial intracellular sodium ion measurement using CoroNa Green. Bio Protoc 7:e2092 42. Guzman LM, Belin D, Carson MJ et al (1995) Tight regulation, modulation, and high-level expression by vectors containing the arabinose PBAD promoter. J Bacteriol 177:4121–4130 43. Gonzalez-Pedrajo B, Minamino T, Kihara M, et al (2006) Interactions between C ring proteins and export apparatus components: a possible mechanism for facilitating type III protein export. Mol Microbiol 60:984–998 44. Che Y-S, Nakamura S, Morimoto YV et al (2014) Load-sensitive coupling of proton translocation and torque generation in the bacterial flagellar motor. Mol Microbiol 91:175– 184 45. Schindelin J, Arganda-Carreras I, Frise E et al (2012) Fiji: an open-source platform for biological-image analysis. Nat Methods 9: 676–682 46. Suzuki Y, Morimoto YV, Oono K et al (2019) Effect of the MotA(M206I) mutation on torque generation and stator assembly in the Salmonella H+-driven flagellar motor. J Bacteriol 201:e00727-18 47. Morimoto YV, Minamino T (2017) Stoichiometry and turnover of the stator and rotor. Methods Mol Biol 1593:203–213

Chapter 9 Purification of the Na+-Driven PomAB Stator Complex and Its Analysis Using ATR-FTIR Spectroscopy Seiji Kojima, Michio Homma, and Hideki Kandori Abstract The flagellar motor of marine Vibrio is driven by the sodium-motive force across the inner membrane. The stator complex, consisting of two membrane proteins PomA and PomB, is responsible for energy conversion in the motor. To understand the coupling of the Na+ flux with torque generation, it is essential to clearly identify the Na+-binding sites and the Na+ flux pathway through the stator channel. Although residues essential for Na+ flux have been identified by using mutational analysis, it has been difficult to observe Na+ binding to the PomAB stator complex. Here we describe a method to monitor the binding of Na+ to purified PomAB stator complex using attenuated total reflectance-Fourier transform infrared (ATR-FTIR) spectroscopy. This method demonstrates that Na+-binding sites are formed by critical aspartic acid and threonine residues located in the transmembrane segments of PomAB. Key words Bacterial flagellum, Motility, Flagellar motor, Stator, PomA, PomB, Decyl maltose neopentyl glycol (DMNG), ATR-FTIR, Sodium binding site

1 1.1

Introduction Background

Motile bacteria can swim in liquid environments or swarm on solid surfaces by rotating a filamentous organ called the bacterial flagellum [1]. This organ consists of three main parts: a long helical filament that generates propulsion to drive the cell body, a rotary motor embedded in the cell surface, and a hook that connects the filament and motor to function as a universal joint (Fig. 1a). Flagellar motor harnesses the ion motive force across the inner membrane as the energy for rotation. Proton (H+) is used as the coupling ion in most bacteria, such as Escherichia coli and Salmonella, but sodium ion (Na+) is used in other bacteria like marine Vibrio [2]. The motor comprises a rotary part (rotor) and multiple stator units that surround the rotor. The stator is a transmembrane protein complex anchored to the peptidoglycan cell wall and conducts the coupling ions through its channel to generate rotational

Tohru Minamino et al. (eds.), Bacterial and Archaeal Motility, Methods in Molecular Biology, vol. 2646, https://doi.org/10.1007/978-1-0716-3060-0_9, © The Author(s), under exclusive license to Springer Science+Business Media, LLC, part of Springer Nature 2023

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Fig. 1 Schematic representation of the Na+-driven polar flagellar motor of Vibrio alginolyticus and its stator proteins. (a) Schematic of the polar flagellar motor. The rotary part of the motor (rotor) comprises mainly the MS-ring and C-ring. The MS ring is connected to the P-ring (attached to the peptidoglycan layer) and L-ring (attached to the outer membrane) via an axial structure called the rod. The hook connects the motor to the filament. Both the hook and filament are covered with a membranous sheath, which is contiguous with the outer membrane. The stator unit is a membrane protein complex consisting of PomA (dark blue) and PomB (light blue). Motor torque is generated by a stator-rotor interaction that couples to the Na+ influx through the stator. The Vibrio motor has additional structures named T-ring and H-ring. (b) Membrane topology of PomA and PomB. PomA has four TM segments with a large cytoplasmic loop, which contains conserved charged residues involved in rotor-stator interaction. Two threonine residues of PomA critical for ion conduction are located in TM3 and TM4. PomB has a single TM segment that contains functionally critical Asp24. The amphipathic plug helix C-terminal to the TM regulates the Na+ influx, and the large periplasmic region containing OmpA-like domain functions to anchor the stator unit to the peptidoglycan layer. The structure of the OmpA-like domain has been solved, and part of it is shown as a ribbon model (PDB id: 3WPW)

force (torque) [3]. The stator is composed of two membrane proteins, MotA and MotB for the H+-driven motor and PomA and PomB for the Na+-driven motor (Fig. 1b). MotA and PomA and MotB and PomB are orthologs, respectively, and they form a hetero-heptamer (A5B2) complex. The A subunit has four transmembrane (TM) segments with a relatively large cytoplasmic loop between its second and third TM segments. This cytoplasmic loop forms the rotor-stator interface and contains conserved charged residues critical for torque generation as well as stator assembly around the rotor. The B subunit has a single TM segment and a large periplasmic region that contains an OmpA-like domain, which is known to bind to the peptidoglycan (PG) layer (Fig. 1b) [4, 5]. Recently high-resolution structures of the stator complexes have been solved by cryo-electron microscopy (cryoEM) image

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analysis [5–7]. This analysis has shown that a ring of five molecules of the A subunit surrounds the TM segments consisting of two B subunit molecules, appearing as if the A subunit ring rotates around the axis of the B subunits anchored to the PG layer (stator rotation model). Mutational analysis has identified many important residues of stator proteins critical for motor rotation [8, 9]. One of these is the critical Asp residue located in the TM segment of the B subunit. It is completely conserved and is believed to be the coupling ion-binding residue in the stator [10]. The binding of either H+ or Na+ to this conserved Asp residue of the B subunit is essential for torque generation, presumably by inducing a conformational change in the A subunit to drive the rotor [11]. Another important element for ion flux through the stator channel is the characteristic amphipathic helix located on the C-terminal side of the TM of the B subunit. This helix regulates ion flux by “plugging” the stator channel and so is called as “plug” segment [12, 13]. In the rotating motor, the active stator units are anchored to the PG layer with their plugs open, allowing the ion influx through the stator channel to couple with rotor-stator interactions. Although residues and regions required for the stator function have been identified by mutational analysis, the mechanism of energy conversion of the flagellar motor remains unknown, mostly due to the lack of in vitro functional studies using purified stator complex. The difficulty with stator purification is attributed to the use of unsuitable detergents to solubilize stator units from membrane. Finally, after myriad concerted efforts on screening detergents over a long time, the optimal detergents have recently been discovered. These are neopentyl glycol detergents (decyl maltose neopentyl glycol (DMNG) or lauryl maltose neopentyl glycol, (LMNG)), which are used for cryoEM single-particle analysis. Solubilization using DMNG allows us to reproducibly obtain highly purified PomAB samples with a sharp single peak on gel-filtration chromatography [14]. LMNG also works well for solubilization of PomAB (manuscript in preparation). This improvement allows the analysis of Na+-binding to the critical Asp residue (Asp24) of PomB. Attenuated total reflectanceFourier transform infrared (ATR-FTIR) spectroscopy is a powerful tool for investigating structural changes in proteins as it detects spectral changes due to changes in intramolecular bonds [15]. ATR-FTIR spectroscopy can use samples in aqueous solution, and so ionic composition can be controlled during measurements. From this point, the Na+-driven PomAB stator complex has a technical advantage over the H+-driven ones because structural changes may be detected by simply changing the Na+ concentration during measurements. Indeed, in 2009, we applied this technique to purified PomAB complex and succeeded in obtaining direct evidence that Na+ binds to the conserved Asp24 residue of PomB [16]. Although we obtained many insightful results from this

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measurement, technical difficulties remained at that time; we needed to purify PomAB from a large amount of cultured Vibrio cells. We used CHAPS (3-[(3-Cholamidopropyl)-Dimethylammonio]-1-Propane Sulfonate]) as the detergent for solubilization, and the purity was not very high. Therefore, a major challenge was to improve the sample preparation methods to facilitate ATR-FTIR analysis of the stator complex. Here, using an improved purification protocol, we can reproducibly obtain much better stator samples for ATR-FTIR analysis, and this has allowed us to improve the quality of FTIR signals and specify the Na+-induced spectral changes [14]. In this chapter, we describe an improved stator protein purification protocol and ATR-FTIR analysis using highly purified samples. From these analyses, we have identified three functionally critical residues in the TM segments of the PomAB stator complex that form the Na+-conducting pathway. 1.2 Overview of the Methods

The first section describes the overproduction and purification of the PomAB stator complex [14, 17]. The stator proteins are overproduced in E. coli from a cold-shock inducible plasmid (pCold I derivative). This allows a rather high level of stator protein expression in the E. coli membrane. We used a hexahistidine tag (His-tag) attached to the C-terminus of PomB, which does not affect the function of the stator. The position of the His-tag is important, because if attached to the C-terminus of PomA, the stator complex will dissociate during purification. The cells are cultured at low temperature (16
C) overnight after cold shock, harvested by centrifugation, and then disrupted in a French press. Membrane fractions are obtained by ultracentrifugation and are stored at 80
C until use. Membrane solubilization can be performed at 30
C by using either DMNG or LMNG. Either detergent is adequate. Solubilized PomAB stator complexes are mixed with a Co-TALON resin for affinity purification. Large aggregates and impurities are still present after affinity purification and are removed by concentration of purified samples using a centrifugal device with a molecular weight cutoff (MWCO) of 100 kDa, followed by size exclusion chromatography (SEC). As shown in Fig. 2, the purified PomAB stator complex shows a sharp single peak on gel filtration analysis (Fig. 2a) and is highly purified as judged by SDS–PAGE stained with Coomassie Brilliant Blue (Fig. 2b) [17]. PomA somehow forms a dimer on SDS-PAGE and does not separate into monomer when treated with the SDS-loading solution. The reason for the formation of the SDS-resistant dimer is unknown. Purified samples can be stored in a refrigerator for up to 1 month, but flash freezing in liquid nitrogen is recommended for long-term storage. In the second section, we describe a method for analyzing Na+induced structural changes in the PomAB stator complex by ATRFTIR spectroscopy. Here, the stator complex is reconstituted into a lipid bilayer and is hydrated during the measurement (Fig. 3a). The

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Fig. 2 Purification of the PomAB stator complex. (a) Typical elution profile of the purified PomAB complex from size exclusion chromatography. The His-tag affinity-purified sample is loaded on an ENrich SEC 650 10  300 column; protein elution is monitored at an absorbance of 280 nm. Black arrow indicates the elution peak. (b) SDS-PAGE gel stained with Coomassie Brilliant Blue. Proteins in peak fractions shown in (a) are analyzed by SDS-PAGE. PomA and PomB are well purified by size exclusion chromatography. For reasons yet unknown, some PomA persists as dimer even after treatment with an SDS-loading solution containing 2-mercaptoethanol. These samples are not boiled. (Figures are partially modified from our previous paper [17])

results reflect the characteristics of the stator complex in its native membrane environment. The measurement is performed initially with a cation-free buffer for background signal measurement, which is then switched to the cation (20 mM) buffer. These measurements are repeated multiple times, and the raw data are averaged to obtain a final spectrum of “20 mM – minus – 0 mM” cation. A previous study has described cation binding FTIR signals for all monovalent cations, not just Na+ [16]. This observation is contrary to the fact that the PomAB stator complex conducts Na+ to generate torque. It is likely that the stator has multiple monovalent cation binding sites, which are either Na+-selective or nonselective. From the aforementioned study, it is difficult to identify the Na+-specific signal because of poor resolution, presumably due to sample impurities. The improved stator protein purification protocol allows us to obtain specifically the Na+-induced FTIR signal. Regarding the “20 mM – minus – 0 mM” Na+ spectrum, strong peaks are observed at the amide-I region (1680–1630 cm1), which are absent in the “20 mM – minus – 0 mM” K+ spectrum (Fig. 3b). Spectra identical to the latter are obtained for 20 mM Li+, Rb+, and Cs+ [14]. Also, strong amide-I bands are absent in the “20 mM – minus – 0 mM” Na+ spectrum of the D24N mutant (Fig. 3b). We are thus able to distinguish Na+-selective and nonselective binding signals, the former of which shows strong amide-I bands. By subtracting nonselective binding signals (“20 mM – minus – 0 mM” K+) from Na+-selective binding signals (“20 mM – minus – 0 mM” Na+), we have obtained the Na+-binding spectrum to the

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Fig. 3 Experimental setup for difference ATR-FTIR spectroscopy and cationinduced difference ATR-FTIR spectra of PomAB. (a) Sample compartment for PomAB reconstituted into lipids. Samples are attached to the ATR cells, which are filled with buffer solution. (b) Difference ATR-FTIR spectra of “20 mM – minus – 0 mM” Na+ (upper spectra) and K+ (upper spectra) for WT and D24N PomAB. In each spectrum, positive and negative bands originate from vibrations in the presence and absence of monovalent cations, respectively. (c) Difference ATR-FTIR spectra of 20 mM Na+ – minus – 20 mM K+, calculated from (b). The difference spectra monitor Na+-binding to the active center of PomAB, whose signals disappear in T158A, T186A, and D24N, but not in D31C

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active center (gray line in Fig. 3c). Strong amide-I bands at 1663 (+)/1649 () cm1 implicate helical structural perturbation upon Na+ binding. The presence of a positive peak at 1115 cm1 is unique to the PomAB stator complex. This complex contains functionally important threonine residues in TM3 (T158) and TM4 (T186), and so the IR bands around the CO stretch region may reflect the signal from these two residues. Measurements of mutant stators with PomA-T158A, PomA-T186A, or PomB-D24N substitution have revealed a complete loss of these specific signals (Fig. 3c). This suggests that Thr158 and Thr186 of PomA, together with Asp24 of PomB, form the Na+ conducting pathway of the PomAB stator complex.

2

Materials

2.1 Purification of the PomAB Stator Complex

All media must be autoclaved. Ampicillin and isopropyl-β-D-thiogalactopyranoside (IPTG) are added to the autoclaved medium after it has cooled down to room temperature. 1. Escherichia coli strain BL21(DE3) harboring a plasmid pCold4-pomAB-His6. This plasmid encodes pomA and pomB-his6 (with or without mutations) under control of the cspA promoter, which is induced in response to cold shock. 2. LB medium: 1% (w/v) Bacto-tryptone, 0.5% (w/v) yeast extract, 0.5% (w/v) NaCl. 3. Ampicillin (100 mg/mL stock in milliQ water, store at 20
C). 4. IPTG (1 M stock in milliQ water, store at 20
C). 5. 2-mercaptoethanol. 6. Na-Pi buffer: 50 200 mM NaCl.

mM

sodium

phosphate,

pH

8.0,

7. DMNG: 10% (w/v) stock in milliQ water, preferably prepared before use but can be stored in the refrigerator. 8. TALON® metal affinity resin. 9. Wash buffer: 50 mM sodium phosphate, pH 8.0, 200 mM NaCl, 20 mM imidazole, 0.05% (w/v) DMNG. 10. Elution buffer: 50 mM sodium phosphate, pH 8.0, 200 mM NaCl, 200 mM imidazole, 0.05% (w/v) DMNG. 11. Amicon® Ultra-4 centrifugal device, with an MWCO of 100 K. 12. ENrich™ SEC 650 10  300 column. 13. Fast protein liquid chromatography system (e.g., AKTA explore system or NGC chromatography system).

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14. K buffer: 20 mM Tris–HCl, pH 8.0, 100 mM KCl, 0.02% (w/v) DMNG. 15. Glass flask (3 L or larger) for cell culture. 16. Open-air orbital shaker for 3 L flask. 17. French press. 18. Tubes for ultracentrifugation. 19. Ultracentrifuge. 20. Open column (plastic or glass). 21. Water bath. 22. Heat block for boiling SDS-PAGE samples. 23. SDS-PAGE system. 24. Spectrophotometer to monitor cell growth. 25. BCA protein assay kit to determine protein concentration. 2.2 ATR-FTIR Analysis of the PomAB Stator Complex

1. 1-palmitoyl-2-oleoyl-sn-phophatidylethanolamine (POPE). 2. 1-palmitoyl-2-oleoyl-sn-phosphatidylglycerol (POPG). 3. Bio-beads SM-2 resin. 4. Perfusion buffer: 10 mM MOPS-Tris [pH 7.0], 5 mM MgCl2. 5. Triple-reflectance Si ATR crystal. 6. N2 gas stream for drying sample lipid. 7. FTIR spectrophotometer equipped with a liquid nitrogencooled MCT detector. 8. Deuterated (D2O) buffer: perfusion buffer prepared using D2O at pD 7.0.

3

Methods

3.1 Purification of the PomAB Stator Complex

1. Inoculate E. coli BL21(DE3) cells harboring pCold4-pomABHis6 from freezer stock into a 50 mL volume of LB medium in a glass flask containing 100 μg/mL ampicillin. Grow the cells overnight at 37
C with shaking at 180 rpm. 2. The next day, dilute 20 mL of the overnight culture in 1.5 L of LB containing 100 μg/mL ampicillin prepared in a 3 L glass flask. We usually prepare two 1.5 L flasks, making a total of 3 L cultures. Grow the cells at 37
C with shaking at 150 rpm. 3. Cell growth is monitored by measuring the absorbance at 660 nm (OD660). When OD660 reaches 0.5, take a small volume of the culture in a 1.5 mL tube to prepare a whole cell sample for SDS-PAGE before chilling the culture (preinduction). Next chill the cell culture in an ice water bath.

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4. After cooling in ice water for 30 min, add IPTG to the culture to a final concentration of 0.5 mM, and then continue the culture overnight at 16
C with shaking at 150 rpm. This cold shock and low temperature induce overproduction of the PomAB stator complex in the E. coli membrane. 5. The next day, again take a small amount of the culture to prepare whole cell samples for SDS-PAGE (post-induction); centrifuge the rest of the culture (7000  g) to collect the cells (see Note 1). 6. Measure the wet weight of the cell pellet. Add 7 mL of ice-cold Na-Pi buffer for 1 g of the cell pellet, and then suspend the cells (see Note 2). 7. Disrupt cells with a French Press (1000 kg/cm2). We usually pass the cell suspension through the French Press twice (see Note 3). To avoid heating the samples, ensure the cell suspension, cylinder, and piston of the French Press are well chilled before the process. 8. Remove unbroken cells by centrifugation (17,000  g for 10 min at 4
C). 9. Subject cell lysates to ultracentrifugation (150,000  g for 30 min at 4
C). Add the same suspension volume of the Na-Pi buffer shown in step 6 to the resultant pellet and homogenize. The samples (membrane suspension) can be stored at 20
C. 10. Next thaw the membrane suspension, and slowly add the DMNG detergent (see Note 4) to the suspension while stirring to a final concentration of 0.5% (w/v). Continue to stir the mixture at 30
C for 30 min, using a water bath to maintain a constant temperature. Dropwise addition of DMNG is recommended following which the turbidity of the suspension is dramatically reduced. 11. Remove insolubilized membrane by ultracentrifugation (150,000  g for 30 min at 4
C). 12. During step 11, pre-equilibrate the TALON metal affinity resin with 2–3 column volumes of Wash buffer. Usually, a column bed volume of 3–4 mL is used for a 3 L culture. 13. Add solubilized membrane samples obtained from step 11 to the pre-equilibrated TALON resin, and gently stir in a cold room at 4
C for 1 h. 14. Load the mixture into an open column, and wash with 3 column volumes of Wash buffer. Note: from this step, all experiments are carried out at room temperature.

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15. Next is elution of the PomAB stator complex by applying 2 column volumes of Elution buffer to the column, and then collect the eluted samples. 16. Confirm the peak fractions by using SDS-PAGE gel stained with Coomassie Brilliant Blue. Samples in peak fractions are collected into a single tube and mixed with 2-mercaptoethanol to a final concentration of 0.1% (v/v) (see Note 5). 17. Perform centrifugation (5000  g for ~10 min at 4
C) to concentrate the sample to approximately 500 μL using the Amicon Ultra-4 centrifugal device (100K MWCO) (see Note 6). 18. Subject the concentrated sample to ultracentrifugation (150,000  g for 10 min at 4
C) to remove any aggregates. 19. Apply the supernatant from step 18 (~500 μL) to the ENrich SEC 650 10  300 column, pre-equilibrated with K buffer using a fast protein liquid chromatography system. 20. Run the column with K buffer at a flow rate of 0.75 mL/min. In this step, Na+ in the samples is removed and replaced with K+. 21. Collect peak fractions and concentrate as in steps 16 and 17 above, without addition of 2-mercaptoethanol. Measure the protein concentration using Pierce BCA method as per manufacturer’s instructions. 22. Store the final purified sample in a refrigerator at 4
C until use (see Note 7). 3.2 ATR-FTIR Analysis of the PomA/ PomB Stator Complex

1. Add purified and detergent-solubilized PomA/PomB stator complex in the K buffer to a lipid mixture of POPE and POPG at a molar ratio of protein complex PomAB: lipid ¼ 1: 40, in a microfuge tube (see Note 8). 2. Prewash the Bio-beads SM-2 with Perfusion buffer, and then add to the mixture; incubate by slow rotation with a rotator. 3. Collect the Bio-beads by centrifugation; the lipid-reconstituted proteins are in the supernatant. Repeat steps 2 and 3 until the supernatant becomes turbid, indicating that the reconstitution of PomAB into the membrane is complete. 4. Using Perfusion buffer, wash the lipid-reconstituted proteins obtained by the procedure above with repeated spin and wash cycles. 5. Take an aliquot of washed sample suspension, and place it on the surface of a triple-reflectance Si ATR crystal. 6. Dry the sample placed on the ATR crystal with a gentle stream of N2 gas.

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7. Rehydrate the dried layer by irrigating with Perfusion buffer at a flow rate of 0.4 mL/min, ensuring the flow cell temperature is maintained at 20
C. 8. Record the ATR-FTIR spectra at 2 cm1 resolution using an FTIR spectrophotometer equipped with a liquid nitrogencooled MCT detector. First, measure a background spectrum (260 interferograms co-added for 5 min) with the sample under a flow of NaCl-free Perfusion buffer. 9. To assess the effect of Na+ on the sample, measure the spectra after switching to a buffer containing 20 mM NaCl. Record the sample spectrum when equilibrated. 10. Repeat the forward/backward (Na+-binding/unbinding) measurements, and then obtain the average of the spectra for 6–30 cycles to produce a raw spectrum as “20 mM – minus – 0 mM” Na+. 11. Calculate the final spectra by subtracting baseline drifts due to protein swelling/shrinkage and free salt/buffer ions in the external buffer from the raw spectra described in step 10.

4

Notes 1. We usually check the expression of the PomAB stator complex by immunoblotting of whole cell samples before induction (preinduction, step 3) and after induction (post-induction, step 5). To save time, typically we disrupt cells immediately after harvesting; we do not store the cell pellet in a deep freezer for later use. 2. Cell concentration in suspension is important for efficient disruption with a French Press. An excessively high concentration of cells usually causes tube clogging in the French Press, making it difficult to control the pressure being applied. 3. Disruption using a sonicator is somewhat inefficient compared to a French Press. In this case, we recommend transforming the E. coli BL21 (DE3) strain with the pLysS plasmid that encodes lysozyme. It digests the peptidoglycan layer and facilitates easy disruption by sonication. Please also see this ref. [17]. 4. LMNG can be used at a similar concentration as DMNG. The purity and stability of the PomAB stator complex solubilized with LMNG are similar to that with DMNG. 5. PomB has three cysteine residues at position 8, 10, and 31. Cys8 and Cys10 are involved in forming an intermolecular disulfide bridge in the two PomB molecules within the stator complex [18]. Usually, 2-mercaptoethanol is added to avoid unnecessary disulfide bridge formation during protein

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concentration steps using the Amicon centrifugal device [14]. However, we recently found that in the absence of 2-mercaptoethanol, no unnecessary disulfide bridges are formed within the stator complex. Therefore, this procedure can be omitted. 6. We perform centrifugation at 5000  g for 5 min and observe the remaining volume in the device. Typically, centrifugation twice for 5 min (total 10 min) is sufficient to achieve a volume of about 500 μL. Centrifugation will take much longer if the buffer contains glycerol. 7. We usually purify the PomAB stator complex immediately before subsequent ATR-FTIR analysis. Thus, it is stored in the refrigerator; purified samples are not flash frozen. Purified PomA/PomB in detergent does not form precipitates for about a month of refrigeration at 4
C. 8. Until the publication of high-resolution stator structures [6, 7], the PomA/PomB complex has been believed to exist as a hetero-hexamer formed by four molecules of PomA and two molecules of PomB, based on results of biochemical studies [19–21]. Consequently, we have used the molecular weight of a hexameric complex to obtain the molar ratio as published earlier [14].

Acknowledgments This research was supported in part by Grants-in-aid for scientific research from the Ministry of Education, Science and Culture of Japan (18K19293 to SK, 24117004 and 23247024 to MH, 15H02391 to HK). We thank Hiroto Iwatsuki and Masayo Iwaki for preparing the figures. References 1. Wadhwa N, Berg HC (2022) Bacterial motility: machinery and mechanisms. Nat Rev Microbiol 20(3):161–173. https://doi.org/ 10.1038/s41579-021-00626-4 2. Terashima H, Kojima S, Homma M (2008) Flagellar motility in bacteria structure and function of flagellar motor. Int Rev Cell Mol Biol 270:39–85. https://doi.org/10.1016/ S1937-6448(08)01402-0 3. Kojima S, Blair DF (2004) The bacterial flagellar motor: structure and function of a complex molecular machine. Int Rev Cytol 233:93–134 4. Takekawa N, Imada K, Homma M (2020) Structure and energy-conversion mechanism of the bacterial Na(+)-driven flagellar motor.

Trends Microbiol 28(9):719–731. https:// doi.org/10.1016/j.tim.2020.03.010 5. Hu H, Santiveri M, Wadhwa N, Berg HC, Erhardt M, Taylor NMI (2022) Structural basis of torque generation in the bi-directional bacterial flagellar motor. Trends Biochem Sci 47(2):160–172. https://doi.org/ 10.1016/j.tibs.2021.06.005 6. Deme JC, Johnson S, Vickery O, Aron A, Monkhouse H, Griffiths T, James RH, Berks BC, Coulton JW, Stansfeld PJ, Lea SM (2020) Structures of the stator complex that drives rotation of the bacterial flagellum. Nat Microbiol 5(12):1553–1564. https://doi.org/10. 1038/s41564-020-0788-8

ATR-FTIR Analysis of Purified PomAB Stator 7. Santiveri M, Roa-Eguiara A, Ku¨hne C, Wadhwa N, Hu H, Berg HC, Erhardt M, Taylor NMI (2020) Structure and function of stator units of the bacterial flagellar motor. Cell 183(1):244–257.e16. https://doi.org/10. 1016/j.cell.2020.08.016 8. Nakamura S, Minamino T (2019) Flagelladriven motility of bacteria. Biomol Ther 9(7): 2 7 9 . h t t p s : // d o i . o r g / 1 0 . 3 3 9 0 / biom9070279 9. Macnab RM (2003) How bacteria assemble flagella. Annu Rev Microbiol 57:77–100 10. Zhou J, Sharp LL, Tang HL, Lloyd SA, Billings S, Braun TF, Blair DF (1998) Function of protonatable residues in the flagellar motor of Escherichia coli: a critical role for Asp 32 of MotB. J Bacteriol 180(10):2729–2735 11. Kojima S, Blair DF (2001) Conformational change in the stator of the bacterial flagellar motor. Biochemistry 40(43):13041–13050 12. Hosking ER, Vogt C, Bakker EP, Manson MD (2006) The Escherichia coli MotAB proton channel unplugged. J Mol Biol 364(5): 921–937 13. Li N, Kojima S, Homma M (2011) Characterization of the periplasmic region of PomB, a Na +-driven flagellar stator protein in Vibrio alginolyticus. J Bacteriol 193(15):3773–3784. https://doi.org/10.1128/JB.00113-11 14. Onoue Y, Iwaki M, Shinobu A, Nishihara Y, Iwatsuki H, Terashima H, Kitao A, Kandori H, Homma M (2019) Essential ion binding residues for Na(+) flow in stator complex of the Vibrio flagellar motor. Sci Rep 9(1):11216. https://doi.org/10.1038/s41598-01946038-6

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15. Kandori H (2020) Structure/function study of photoreceptive proteins by FTIR spectroscopy. Bull Chem Soc Jpn 93(7):904–926. https:// doi.org/10.1246/bcsj.20200109 16. Sudo Y, Kitade Y, Furutani Y, Kojima M, Kojima S, Homma M, Kandori H (2009) Interaction between Na+ ion and carboxylates of the PomA-PomB stator unit studied by ATR-FTIR spectroscopy. Biochemistry 48(49):11699–11705. https://doi.org/10. 1021/bi901517n 17. Nishikino T, Iwatsuki H, Mino T, Kojima S, Homma M (2020) Characterization of PomA periplasmic loop and sodium ion entering in stator complex of sodium-driven flagellar motor. J Biochem 167(4):389–398. https:// doi.org/10.1093/jb/mvz102 18. Yorimitsu T, Kojima M, Yakushi T, Homma M (2004) Multimeric structure of the PomA/ PomB channel complex in the Na+-driven flagellar motor of Vibrio alginolyticus. J Biochem 135(1):43–51 19. Braun TF, Al-Mawsawi LQ, Kojima S, Blair DF (2004) Arrangement of core membrane segments in the MotA/MotB proton-channel complex of Escherichia coli. Biochemistry 43(1):35–45 20. Kojima S, Blair DF (2004) Solubilization and purification of the MotA/MotB complex of Escherichia coli. Biochemistry 43(1):26–34 21. Sato K, Homma M (2000) Functional reconstitution of the Na(+)-driven polar flagellar motor component of vibrio alginolyticus. J Biol Chem 275(8):5718–5722

Chapter 10 Purification of Na+-Driven MotPS Stator Complexes and Single-Molecule Imaging by High-Speed Atomic Force Microscopy Naoya Terahara and Noriyuki Kodera Abstract The stator unit of the bacterial flagellar motor coordinates the number of active stators in the motor by sensing changes in external load and ion motive force across the cytoplasmic membrane. The structural dynamics of the stator unit at the single-molecule level is key to understanding the sensing mechanism and motor assembly. High-speed atomic force microscopy (HS-AFM) is a powerful tool for directly observing dynamically acting biological molecules with high spatiotemporal resolution without interfering with their function. Here, we describe protocols for single-molecule imaging of the Na+-driven MotPS stator complex by HS-AFM. Key words Bacterial flagellar motor, Membrane protein, Ion channel, Atomic force microscopy, Single-molecule measurement

1

Introduction Many motile bacteria can move toward favorable or away from unfavorable environments by rotating long filamentous structures called flagella [1] (Fig. 1a). The flagellar motors of Escherichia coli and Salmonella enterica utilize the proton (H+) motive force for torque generation, while those of Vibrio spp. and a particular group of Bacillus spp. utilize the sodium ion (Na+) motive force [2, 3]. The motor structure is conserved across bacterial species and strains and consists of a rotor and a dozen stator units [4] (Fig. 1b). The stator unit functions as a transmembrane ion channel, and the torque is generated by the interaction between the rotor and stator unit that occurs when the coupling ions flow through the stator channel [2]. The stators are classified into three groups based on their primary sequence similarity and coupling ions for flagella rotation. Many bacteria have H+-driven MotAB as the stator unit, while marine Vibrio spp. and alkalophilic

Tohru Minamino et al. (eds.), Bacterial and Archaeal Motility, Methods in Molecular Biology, vol. 2646, https://doi.org/10.1007/978-1-0716-3060-0_10, © The Author(s), under exclusive license to Springer Science+Business Media, LLC, part of Springer Nature 2023

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Fig. 1 Bacterial flagellar motor of Bacillus subtilis. (a) Electron micrograph of the B. subtilis BR151MA cell. (b) Schematic diagram of the Bacillus flagellar motor. (c) Na+-dependent assembly and disassembly model of the MotPS complex. An increase in the Na+ concentration increases the number of active MotPS stator units around the rotor, turning it into a hybrid motor

Bacillus spp. have Na+-driven PomAB and MotPS as the stator unit, respectively [3]. Both stator types appear to have been optimized to harness specific ions according to the environmental conditions. Neutralophilic Bacillus subtilis has Na+-driven MotPS in addition to H+-driven MotAB [5]. The number of active MotPS stator units increases with elevated external Na+ concentrations, so the motor becomes a hybrid complex consisting of both the MotAB and the MotPS stator units around the rotor [6] (Fig. 1c). Furthermore, the stator units quickly associate with or dissociate from the rotor in response to changes in the external Na+ concentration, so that the motor maintains optimal performance for a variety of environmental conditions [7]. These observations suggest that MotPS may have Na+-dependent assembly–disassembly dynamics of the rotorstator complex. This raises a possibility that MotPS acts as a Na+ sensor as well. Recently, high-resolution cryoEM image analyses revealed that the H+-driven MotAB complex consists of five copies of MotA and two copies of MotB with two distinct H+ pathways [8, 9]. The asymmetric nature of the 5:2 subunit stoichiometry proposes a plausible model that the MotA pentamer ring may rotate around the MotB dimer as a central stalk. However, to understand the mechanism underlying the biological function of the stator

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complex, simultaneous evaluation of the structure and dynamics is necessary. Such simultaneous evaluation has recently become possible by the advent of the high-speed atomic force microscopy (HS-AFM) imaging technology. For example, HS-AFM has directly visualized two-headed myosin V walking along actin filaments, bacteriorhodopsin responding to light, and the rotational propagation of conformational changes in the α3β3 subcomplex of the F1-ATPase. Such dynamic HS-AFM images have provided mechanistic insights into their molecular processes [10– 12]. Recently, we have elucidated the Na+-sensing mechanism of the MotPS stator complex by HS-AFM [13]. Here, we describe the protocols for purifying the MotPS complex, its HS-AFM imaging, and image analysis.

2

Materials Prepare all solutions using analytical grade regents and ultrapure water. All buffers used for HS-AFM imaging should be filtered by 0.22 μm syringe-driven filters. Store all reagents at 4 °C unless otherwise specified.

2.1 Reagents and Buffers Used for Preparation of Membrane Fractions

1. Expression plasmid: pET21b vector encoding His6-tagged MotP/MotS (see Note 1). 2. Expression strain: Escherichia coli BL21 (DE3) (see Note 2). 3. LB medium: 1% (w/v) Bacto tryptone, 0.5% (w/v) yeast extract, 0.5% (w/v) NaCl. 4. LBG medium: L-broth with 0.1% (w/v) glucose. 5. LB plate: LB medium with 1.5% (w/v) Bacto agar. Ampicillin is added to the medium at a final concentration of 50 μg/mL. 6. Isopropyl-β-D-1-thiogalactopyranoside (IPTG): 0.6 M aqueous solution, sterilized with a 0.22 μm syringe-driven filter. 7. Tris–HCl buffer: 50 mM Tris–HCl, pH 8.0. 8. Extraction buffer: 50 mM Tris–HCl, pH 8.0, 50 μg/mL DNase I, cOmplete protease inhibitor cocktail (1 tablet/50 mL buffer). 9. Membrane buffer: 50 mM Tris–HCl, pH 8.0, 10% (v/v) glycerol. 10. 5 L Erlenmeyer flask with baffles. 11. Autoclave. 12. Incubator shaker. 13. Spectrophotometer. 14. Floor-standing centrifuge (e.g., Beckman Coulter, model: Avanti HP-26XP).

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15. Centrifuge bottles (e.g., Beckman Coulter, model: 393033). 16. French pressure cell press (e.g., Central Scientific Commerce, model: FA-078A). 17. French pressure cell (e.g., Central Scientific Commerce, model: FA-032). 18. Refrigerated microcentrifuge. 19. Centrifuge tubes. 20. Ultracentrifuge (e.g., Beckman Coulter, model: Optima XL-80). 21. Ultracentrifuge tubes (e.g., Beckman Coulter, model: 355631). 2.2 Reagents and Buffers Used for Purification of MotPS Complexes

1. Decyl maltose neopentyl glycol (DMNG): 10% (w/v) solution in water (prepare just before use). 2. Solubilization buffer: 50 mM Tris–HCl, pH 8.0, 10% (v/v) glycerol, 500 mM NaCl, 1% (w/v) DMNG (see Note 3). 3. Ni-NTA agarose resin. 4. Disposable 5 mL column for gravity-flow procedures. 5. Wash buffer: 50 mM Tris–HCl, pH 8.0, 5% (v/v) glycerol, 500 mM NaCl, 50 mM imidazole, 0.1% (w/v) DMNG. 6. Elution buffer: 50 mM Tris–HCl, pH 8.0, 500 mM NaCl, 400 mM imidazole, 0.05% (w/v) DMNG. 7. SEC1 buffer: 20 mM Tris–HCl, pH 8.0, 500 mM NaCl, 0.05% (w/v) DMNG. 8. SEC2 buffer: 20 mM Tris–HCl, pH 8.0, 150 mM NaCl. 9. Amphipols A8-35: 5% (w/v) solution in water (prepare just before use). 10. Bio-Beads SM-2 Adsorbents: Rinse the required amount of beads with methanol and then with a detergent-free buffer before use. Do not allow the beads to dry. 11. Mini-disk rotator (e.g., Bio-Craft, model: BC-710). 12. Fast protein liquid chromatography system (e.g., GE Healthcare, AKTA purification system). 13. Size exclusion chromatography column (e.g., GE Healthcare, Superose 6 10/300 GL and Superdex 200 10/300 GL). 14. Spectrophotometer.

2.3

HS-AFM Imaging

1. Observation buffer: 20 mM Tris–HCl, pH 8.0 with or without various concentrations of NaCl or KCl. 2. NaCl buffer: 20 mM Tris–HCl, pH 8.0, 150 mM NaCl. 3. KCl buffer: 20 mM Tris–HCl, pH 8.0, 150 mM KCl.

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4. NiCl2 solution: 1 mM NiCl2. 5. CoroNa Green Sodium Indicator. 6. Sodium Standard Solution Na1000. 7. Fluorescence spectrophotometer. 8. HS-AFM apparatus (an apparatus with the same performance is commercially available from Research Institute of Biomolecule Metrology, model: SS-NEX) [14, 15]. 9. Vibration isolation table (e.g., Herz, model: TDI-107100LA). 10. Cantilever (Olympus, model: BL-AC10DS-A2) (see Note 4). 11. Glass rod (Japan Cell, model: 2.0φ × 2 mm height). 12. Clear Mica disk (Fuuchi Chemical, model: clear-mica substrate 1.5φ × 0.1 mm thickness). 13. Software for AFM image analysis (a laboratory-built software, model: Kodec 4.4.7.39) (see Note 5). 14. XYZ-axis manual manipulator (e.g., Narishige, model: M-152). 15. High-precision syringe pump system (Harvard, model: 11 Pico Plus Elite) (see Note 6).

3

Methods

3.1 Preparation of Membrane Fractions Containing His6Tagged MotPS Complexes

1. As a preculture, inoculate the transformed cells on an LB plate, and incubate at 30 °C for approximately 12–14 h. 2. The whole bacterial lawn on the surface of the LB plate is collected by using a sterile inoculation loop and inoculate into 2 L of LBG medium in a 5 L Erlenmeyer flask with baffles (see Note 7). To allow aerobic conditions for growth, incubate the flask in an orbital shaker at 30 °C with shaking at 100 rpm. 3. When the optical density of the liquid culture at 600 nm reaches between 0.8 and 1.2, add IPTG to a final concentration of 0.6 mM to induce protein expression. Incubate the culture at 24 °C with shaking at 100 rpm for 4 h (see Note 8). 4. Lower the temperature of the shaker to 18 °C, and shake the culture at 80 rpm for 4 h (see Note 8). 5. Centrifuge the culture at 10,000 × g for 10 min at 4 °C to harvest the cells. 6. Wash the cell pellet with 100 mL of Tris–HCl buffer. 7. Centrifuge the suspension at 10,000 × g for 10 min at 4 °C to harvest the cells. 8. Resuspend the cell pellet in 50 mL of extraction buffer.

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9. Disrupt the cells by passage through a French pressure cell (see Note 9). 10. Centrifuge the cell lysate at 20,000 × g for 10 min at 4 °C to remove undisrupted cells and debris. 11. Transfer the supernatant to a high-speed centrifuge tube, and ultracentrifuge at 110,000 × g for 60 min at 4 °C to pellet the membrane fragments and vesicles. 12. Discard the supernatant, and suspend the membrane pellet in 5 mL of membrane buffer. 13. Determine the protein concentration by the Lowry method, and adjust the volume using membrane buffer so that the final concentration is 10 mg/mL (see Note 10). 14. Store at -20 °C prior to purification. 3.2 Purification of His6-Tagged MotPS Complexes

1. Dilute the membrane fraction in solubilization buffer to a final concentration of approximately 2 mg/mL in order to solubilize membrane proteins. Incubate in an end-over-end rotator at 4 °C for 30 min. 2. Ultracentrifuge the sample at 110,000 × g for 30 min at 4 °C to remove insoluble debris, and transfer the supernatant containing the solubilized fraction to a centrifuge tube at 4 °C. 3. A 10 mL Ni-NTA resin slurry is used for the 10 mL solubilized fraction. Add an appropriate amount of the resin into a disposable column. 4. Equilibrate the Ni-NTA resin with 5× volumes of wash buffer. 5. Add the equilibrated Ni-NTA resin to the solubilized fraction, and mix well using an end-over-end rotator at 4 °C for 30 min. 6. Add the MotPS-bound Ni-NTA resin into the same column, and allow it to drain by gravity flow. 7. Wash the column with 5× volumes of wash buffer to remove the unbound proteins. 8. Elute the His6-tagged MotPS complexes in 1× volume of elution buffer. 9. Analyze the eluted fraction by western blotting using anti-Histag antibody to check if the fraction contains the target protein (see Note 11). 10. Load the elution fraction onto a Superose 6 column equilibrated with SEC1 buffer (Fig. 2b). 11. To replace DMNG with amphipols (Fig. 2a), mix the eluted peak fractions correspond with the MotPS complex with amphipols A8-35 at a 1–3 weight ratio using an end-over-end rotator at 4 °C for 12 h (see Note 12).

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Fig. 2 Structural characterization and purification by size exclusion chromatography (SEC). (a) Sample preparation for HS-AFM imaging following membrane fractions preparation. Replace the detergent with amphipols to reduce the influence of the detergent, and set the ionic strength freely. (b) SEC elution profiles (left panel) of solubilized His6-MotP/MotS by decyl-β-D-maltoside (DM, red), dodecyl-β-D-maltoside (DDM, orange), and decyl maltose neopentyl glycol (DMNG, blue). V0 indicates the position of void volume. A standard curve is made from the elution volume of main peaks of ferritin (440 kDa), catalase (232 kDa), and ovalbumin (44 kDa). DMNG is most effective for solubilizing MotPS in the complex form. SEC elution profiles (right panel) of amphipol-treated His6-MotP/ MotS (orange) and MotP/MotS-His6 (blue). The profiles should be the same even if the position of the His-tag in the complex changes

12. Add the activated Bio-Beads at a final concentration of 15 mg/ mL to remove DMNG, and incubate in an end-over-end rotator at 4 °C for 48 h (see Note 13). 13. Ultracentrifuge the supernatant at 110,000 × g for 30 min at 4 °C to remove the Bio-Beads and insoluble debris.

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14. Load the supernatant onto a Superdex 200 column equilibrated with SEC2 buffer (Fig. 2b). 15. Analyze the eluted peak fractions containing the MotPS complex by SDS-PAGE and CBB staining to check the purity of the target proteins in the fraction (see Note 11). 16. Measure the absorbance at 280 nm of the fractions by spectrophotometer to estimate the concentration of the MotPS complex in the fraction. 3.3

HS-AFM Imaging

1. Set the cantilever in the cantilever holder. 2. Mount the cantilever holder onto the HS-AFM apparatus. 3. Wash the liquid cell with 100 μL each of 3 M KCl solution and Milli-Q water to remove any dust. 4. Fill the liquid cell with 60 μL of the observation buffer. 5. Align the cantilever position so that laser reflected from the cantilever is centered on the two-segmented photodetector and its reflected intensity is maximized. 6. Measure the power spectrum of thermal fluctuations in the cantilever deflection using an oscilloscope equipped with a real-time FFT function to determine the resonant frequency of cantilever. 7. Excite the cantilever around the resonant frequency by applying AC voltage to the piezo actuator attached to the cantilever holder. Set the cantilever’s free oscillation amplitude (A0) to 0.5 V, which corresponds to the peak-to-peak amplitude (2A0) of 3–4 nm. 8. Clean the surface of the z-piezo (z-scanner) with acetone to remove any dust. 9. A glass rod with a thin mica disk glued to the top by epoxy is used as a sample stage (Fig. 3a). Glue the sample stage onto the top of the z-scanner using a drop of nail polish, and wait at least for 10 min for the nail polish to cure. 10. Prepare a freshly cleaved mica surface by removing the top layers of mica using scotch tape, and check the peeled mica by eye whether the mica surface is evenly cleaved. 11. Deposit 2 μL of 1 mM NiCl2 on the mica surface, and incubate it for 3 min. Wash the mica surface thoroughly with 20 μL of Milli-Q water (Fig. 3a) (see Note 14). 12. Dilute the protein sample (final concentration approximately 10 nM) in the observation buffer, and deposit 2 μL of the sample onto the mica surface. Incubate it for 3 min at room temperature. Wash the mica surface with 20 μL of the observation buffer to remove unattached particles.

Fig. 3 Single-molecule imaging of the MotPS complex by HS-AFM. (a) Sample preparation on the sample stage. Place a sample solution on the freshly cleaved mica disk surface for 3 min, and rinse using an observation buffer solution and a piece of Kimwipe cleaning paper while avoiding drying the surface. (b) Schematic diagram of the MotPS complex, which is predicted to consist of five copies of MotP and two copies of MotS (left panel). The C-terminal of the MotS subunit (MotSC) forms a dimer through an interaction between its peptidoglycan-binding domains. The stator complex is visualized by HS-AFM imaging as two ellipsoid domains connected by a flexible linker (right panel). The large and small ellipsoid domains correspond with the transmembrane (TM) domain and MotSC, respectively. Color bar shows a range of particle height. Scale bar indicates 10 nm in the image. (c) Measurement of the height and the center-to-center distance between the small and large domains. Sequential HS-AFM image of a MotPS molecule is recorded at 200 ms/frame (upper panel). Scale bars show 10 nm. The cross-section profile along the line from the filled arrow to the open arrow in each image is shown as a red line (lower panel). To measure the peak height and peak-to-peak distance, the profile is fitted by two Gaussian functions as indicated by blue line

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13. Immerse the sample stage in the liquid cell filled with the observation buffer. 14. While monitoring the position of the cantilever and the sample stage with a digital camera, manually adjust the position of the sample stage surface so that the cantilever is located in a mica disk of the sample stage surface. 15. Set the feedback set-point amplitude (As) to 0.45 V which corresponds to the peak-to-peak amplitude (2As) of 2.7–3.6 nm, and switch on the PID feedback controller. Under these conditions, the average tapping force can be approximated as 20–30 pN using the following equation: pffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffi kc A 0 2 - A s 2: hF i = 2Q c

16. Start the tip-sample approach slowly by driving a stepping motor attached to the scanner. 17. Once the cantilever tip makes contact with the sample, retract the stepping motor so that the output voltage of the PID feedback controller approaches 0 V. Modulate the integral and proportional gain of the PID feedback controller to visualize the surface structure. 18. Start imaging and adjust As so that clear images are obtained. 19. Find the MotPS molecules by moving stage in the xy-plane to examine different areas (Fig. 3b). 3.4 Buffer Exchanging System

1. Degas NaCl buffer and KCl buffer with an aspirator for 30 min to remove bubbles and load them into each infusion syringe. 2. Arrange the tip ends connected to the infusion syringes into the liquid cell by using xyz-axis manual manipulator (Fig. 4a). 3. To measure precise Na+ concentrations of the buffer, fractionate the buffer in the liquid cell sequentially during running the pump system. 4. Add the CoroNa Green Na+-sensitive fluorescent dye into the fractions to a final concentration of 10 μM, and measure the fluorescence intensity (λex = 485 nm, λem = 516 nm) using a fluorescence spectrophotometer (see Note 15). To obtain a calibration curve of the Na+ concentration, measure the fluorescence intensity of the CoroNa Green over a wide range of Na+ concentration (0–150 mM) using a sodium standard solution. This allows calibration of the operating time and flow rate of the pump system and the Na+ concentration of the buffer in the cell (Fig. 4b). 5. Start the pump system during HS-AFM imaging, and capture the dynamic structural changes of the target protein (Fig. 4c).

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Fig. 4 Single-molecule imaging of MotSC during the structural transition triggered by Na+. (a) Schematic diagram of the buffer exchange system. A constant flow pump simultaneously injects the buffer into one side and withdraws from the other side of the liquid cell at the flow rate of 2.5 μL/s. This system gradually changes the composition of the buffer in the liquid cell without changing the amount of the liquid in the cell. By connecting two injection syringes, it is possible to gradually increase the NaCl (left panel) or KCl (right panel) concentration in the buffer. (b) An estimated NaCl concentration (blue curve) and actual Na+ concentration (green triangle) in the buffer when the salt is gradually exchanged from 150 mM KCl to 150 mM NaCl (left panel) or from 150 mM NaCl to 150 mM KCl (right panel). The mean and standard deviation are calculated from three independent measurements. (c) Real-time imaging of MotPS by exchanging the salt in the buffer from 150 mM KCl to 150 mM NaCl (upper panel) or from 150 mM NaCl to 150 mM KCl (lower panel). MotSC becomes suddenly folded after approximately 60 s (upper second and third panels) from the buffer exchange indicated as 0.00 s (150 mM KCl) and unfolded after approximately 5 s (lower second and third panels) from the buffer exchange indicated as 0.00 s (150 mM NaCl). These results indicate that an increase in the Na+ concentration induces the conformational transition in the MotSC, making the MotPS complex an active stator unit in the presence of Na+. All the images are recorded at 250 ms/frame. Scale bar indicates 10 nm

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Image Analysis

1. Pretreat HS-AFM images with a low-pass filter to remove spike noise and a flattening filter to make the overall xy-plane flat, using a laboratory-built software. 2. Take cross-section profiles of the imaged two ellipsoidal structures along their centers of gravity. Extract and list the peak heights of each ellipsoid in each frame of interest (see Note 16). 3. Subtract the average height of the sample-free substrate surface from the peak height of the cross-section profiles (Fig. 3c). 4. Make a histogram of the background-subtracted peak heights, and perform Gaussian fitting to determine their distribution. 5. Measure the center-to-center distance. 6. A minimum of 20 individual molecules analyzed over 600 total frames are required for a robust analysis.

4

Notes 1. Stator protein is purified by affinity chromatography using hexa-histidine tag (His6-tag) (Fig. 2a). The expression vectors are constructed by cloning the full-length motP and motS genes from B. subtilis into a pET21b vector as previously described [13]. The His6-tag should be introduced to either the N-terminus of the MotP (His6-MotP/MotS) or the C-terminus of the MotS (MotP/S-His6); occlusion of the C-terminus of the MotP or the N-terminus of the MotS blocks the rotation of the flagellar motor. A His6-tag seems to be suitable for protein purification and stable adhesion to the mica surface for HS-AFM imaging [16]. 2. First, the conditions for the expression of the membrane protein should be optimized. The E. coli C41 (DE3) and C43 (DE3) strains are also suitable for expressing membrane proteins, but BL21 (DE3) is most effective for expressing the MotPS. Furthermore, individual colonies may differ significantly in their expression levels even if transformed with the identical plasmid. Therefore, it is necessary to select several colonies from each transformation. Samples taken under various conditions can be analyzed, and their MotPS expression levels determined by western blotting using the His6-tag included in the construct. Pick up the highly expressed clones, prepare their respective 20% (v/v) glycerol stocks, and store at -80 °C. 3. The difficulties in the purification of membrane protein complexes originate from their properties as membrane proteins. They are very hydrophobic and may have single or several transmembrane segments; many of them also assemble into multi-subunit complexes. The choice and concentration of detergents is one of the key factors to the purification of intact

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membrane protein complexes. However, there is no universally applicable detergent for all membrane proteins, so careful screening is needed. Detergents are classified according to their structure and fall into three major categories: nonionic [octyl glucoside (OG), decyl maltoside (DM), dodecyl maltoside (DDM), decyl maltose neopentyl glycol (DMNG), cyclohexyl maltoside (CYMAL), tetraethylene glycol monooctyl ether (C8E4), Digitonin, etc.], ionic (sodium cholate, etc.), and zwitterionic [cholamidopropyl dimethylammonio propane sulfonate (CHAPS), lauryldimethylamine oxide (LDAO), and phosphocholine (Fos-Choline), etc.]. We have examined the effects of nearly all detergents on the solubilization and maintenance of the MotPS complex by size exclusion chromatography (Fig. 2b). Most of these detergents destabilize the complex, and DMNG is most effective for solubilizing the intact complex. 4. The cantilevers used are BL-AC10DS-A2 with a resonant frequency ~0.5 MHz, quality factor (Qc) ~1.5 in water and a spring constant (kc) ~0.1 N/m. A probe tip is grown on the original tip end of the cantilever by electron beam deposition (EBD) and further sharpened by argon plasma etching. The detailed procedures for preparing the sharp EBD tips and cleaning cantilevers used are described elsewhere [17]. 5. This software is available at https://elifesciences.org/con tent/4/e04806/ar ticle-data#fig-data-supplementar ymaterial [18]. 6. The buffer in the liquid cell can be exchanged during HS-AFM imaging by installing a high-precision syringe pump system with minor modifications to the liquid cell of the HS-AFM apparatus. This enables the visualization of dynamic protein structural changes caused by a specific substance [18]. We use a constant-pressure and constant-flow pump system with 10 mL syringes that simultaneously infuses and withdraws buffer from opposite sides of the liquid cell. The flow rate can be adjusted from 0.2 nL/s to 0.2 mL/s, allowing the buffer composition to be gradually exchanged. In the case of the MotPS complex, the pump is operated at a flow rate of 2.5 μL/s, and the buffer volume in the liquid cell is kept constant at approximately 60 μL. Details of this pump system setup are described elsewhere [19]. 7. From the screening results, the MotPS expression level of the cells precultured on the LB plate is higher than that of the cells precultured in the liquid LB medium. 8. The expression level is higher at 30 °C than at 37 °C. In general, the amount of inclusion body tends to decrease as the culture temperature and shaking speed are gradually lowered.

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9. A French pressure cell is ideal because it generates less heat. But it is also possible to disrupt the cells using a sonicator where sonicate the cell lysate for two rounds of 10 min each while keeping it on ice. 10. Several protein assays are available, but the Lowry method is recommended for membrane proteins [20]. Prepare a Lowry solution by mixing a reagent A [2.0% (w/v) Na2CO3 in 0.1 N NaOH] and a reagent B [0.5% (w/v) CuSO4·5H2O in 1.0% (w/v) sodium citrate] at a 50 to 1 volume ratio just before use. Dilute 5 μL of the membrane fraction to 100 μL with the membrane buffer. Remember to prepare 100 μL of each concentration of BSA standard (range of 50–800 μg/mL) to absolutely quantify the amount of total protein. Add 800 μL of Lowry solution, and incubate for 10 min at room temperature. Add 100 μL of a Folin phenol reagent, and incubate for 30 min at room temperature. Measure the absorbance at 750 nm of the samples by a spectrophotometer, and prepare a standard curve to determine the protein concentration in the fraction. 11. Sodium dodecyl sulfate-polyacrylamide gel electrophoresis (SDS-PAGE) and western blotting are performed using standard protocols. MotP and MotS in the elution fractions are separated using 12.5% SDS-polyacrylamide gels. Since the molecular weights of MotP (27 kDa) and MotS (24 kDa) are very similar, it is somewhat difficult to distinguish the two proteins by Coomassie brilliant blue (CBB) staining following electrophoresis. Therefore, Tris-Tricine buffer, which is more suitable for separating low molecular weight proteins than Tris-Glycine buffer, is recommended as the SDS-PAGE running buffer [21]. Following electrophoresis, proteins separated on the polyacrylamide gel are electrically transferred to a nitrocellulose membrane. After blocking the membrane with 10% (w/v) skim milk in TBST [Tris-buffered saline containing 0.05% (v/v) Tween-20], incubate in TBST containing polyclonal anti-MotP, anti-MotS, and anti-His6-tag antibodies as previously described [6]. Detection is performed by reacting HRP-conjugated antibody as a secondary antibody followed by addition of ECL substrate. 12. The properties of the detergent depend on the concentration, ionic strength, and reaction temperature. In general, micelle size is larger in a solution with high ionic strength than in a solution with low ionic strength. Further, the critical micelle concentration (CMC) of the nonionic detergent decreases with increasing the temperature. The NaCl or KCl concentration in the observation buffer is changed during HS-AFM imaging. Replace the detergent with amphipols to avoid the effects by the salt concentration change.

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13. The reaction time of amphipols and the detergent removal time by Bio-Beads seem to vary depending on the CMC of the detergent. In the case of DMNG with a low CMC, it takes a very long time to remove DMNG. Increasing the amount of Bio-Beads to reduce the detergent removal time is not recommended because this will promote protein aggregation. 14. His6-tagged MotPS complexes can be easily immobilized through metal-chelate affinity by treating the bare mica surface with Ni2+. 15. It is also possible to use a Na+ meter (HORIBA, model: LAQUAtwin-Na) instead of a fluorescence spectrophotometer to measure the Na+ concentration. 16. Note that the width of the object imaged by AFM is larger than its actual width because of a convolution effect caused by the tip diameter. In contrast, the height of the object can be accurately measured [22]. Therefore, the peak profile representing these domains can be used to accurately measure the distance between the two domains as described previously [23].

Acknowledgments We thank Prof. Toshio Ando and Dr. Steven J. McArthur for critical reading and the English language improvement of the paper. This research has been supported by JSPS KAKENHI Grant 21K06073 to N.T. and 20H00327 to N.K. References 1. Miyata M, Robinson RC, Uyeda TQP et al (2020) Tree of motility – a proposed history of motility systems in the tree of life. Genes Cells 25:6–21 2. Berg HC (2003) The rotary motor of bacterial flagella. Annu Rev Biochem 72:19–54 3. Minamino T, Terahara N, Kojima S et al (2018) Autonomous control mechanism of stator assembly in the bacterial flagellar motor in response to changes in the environment. Mol Microbiol 109:723–734 4. Minamino T, Imada K (2015) The bacterial flagellar motor and its structural diversity. Trends Microbiol 23:267–274 5. Ito M, Hicks DB, Henkin TM et al (2004) MotPS is the stator-force generator for motility of alkaliphilic Bacillus, and its homolog is a second functional Mot in Bacillus subtilis. Mol Microbiol 53:1035–1049

6. Terahara N, Noguchi Y, Nakamura S et al (2017) Load- and polysaccharide-dependent activation of the Na+-type MotPS stator in the Bacillus subtilis flagellar motor. Sci Rep 7: 46081 7. Terahara N, Namba K, Minamino T (2020) Dynamic exchange of two types of stator units in Bacillus subtilis flagellar motor in response to environmental changes. Comput Struct Biotechnol J 18:2897–2907 8. Santiveri M, Roa-Eguiara A, Ku¨hne C et al (2020) Structure and function of stator units of the bacterial flagellar motor. Cell 183:244– 257 9. Deme JC, Johnson S, Vickery O et al (2020) Structures of the stator complex that drives rotation of the bacterial flagellum. Nat Microbiol 5:1553–1564

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10. Kodera N, Yamamoto D, Ishikawa R et al (2010) Video imaging of walking myosin V by high-speed atomic force microscopy. Nature 468:72–76 11. Shibata M, Yamashita H, Uchihashi T et al (2010) High-speed atomic force microscopy shows dynamic molecular processes in photoactivated bacteriorhodopsin. Nat Nanotechnol 5:208–212 12. Uchihashi T, Iino R, Ando T et al (2011) High-speed atomic force microscopy reveals rotary catalysis of rotorless F1-ATPase. Science 333:755–758 13. Terahara N, Kodera N, Uchihashi T et al (2017) Na+-induced structural transition of MotPS stator assembly of the Bacillus flagellar motor. Sci Adv 3:eaao4119 14. Ando T, Kodera N, Takai E et al (2001) A high-speed atomic force microscope for studying biological macromolecules. Proc Natl Acad Sci U S A 98:12468–12472 15. Ando T, Uchihashi T, Fukuma T (2008) Highspeed atomic force microscopy for nanovisualization of dynamic biomolecular processes. Prog Surf Sci 83:337–437 16. Sumino A, Uchihashi T, Oiki S (2017) Oriented reconstitution of the full-length KcsA potassium channel in a lipid bilayer for AFM imaging. J Phys Chem Lett 8:785–793 17. Uchihashi T, Kodera N, Ando T (2012) Guide to video recording of structure dynamics and

dynamic processes of proteins by high-speed atomic force microscopy. Nat Protoc 7:1193– 1206 18. Ngo KX, Kodera N, Katayama E et al (2015) Cofilin-induced unidirectional cooperative conformational changes in actin filaments revealed by high-speed atomic force microscopy. eLife 4:04806 19. Miyagi A, Chipot C, Rangl M et al (2016) High-speed atomic force microscopy shows that annexin V stabilizes membranes on the second timescale. Nat Nanotechnol 11:783– 790 20. Lowry OH, Rosebrough NJ, Farr AL et al (1951) Protein measurement with the Folin phenol reagent. J Biol Chem 193:265–275 21. Sch€agger H, Jagow GV (1987) Tricine-sodium dodecyl sulfate-polyacrylamide gel electrophoresis for the separation of proteins in the range from 1 to 100 kDa. Anal Biochem 166:368– 379 22. Kuznetsov YG, McPherson A (2011) Atomic force microscopy in imaging of viruses and virus-infected cells. Microbiol Mol Biol Rev 75:268–285 23. Kodera N, Uchida K, Ando T et al (2015) Two-ball structure of the flagellar hook-length control protein FliK as revealed by high-speed atomic force microscopy. J Mol Biol 427:406– 414

Chapter 11 High-Resolution Rotation Assay of the Bacterial Flagellar Motor Near Zero Loads Using a Mutant Having a Rod-Like Straight Hook Shuichi Nakamura and Tohru Minamino Abstract The bacterial flagellar motor is embedded within the cell envelop and rotates the long helical filament, which acts as a molecular screw to propel the bacterium. The flagellar motor comprises a rotor and a dozen stator units, converting ion flux through the stator unit into torque. However, the energy coupling mechanism has not been fully understood. Various methods for rotation measurement have been developed to understand the rotation mechanism of the flagellar motor, but the most preferred method in recent studies is a bead assay, which tracks the rotation of a micron to submicron bead attached to the partially sheared flagellar filament at high temporal and spatial resolutions. The bead assay allows us to assess the motor rotation over a wide range of external load, but the elasticity of the axial parts of the flagellum, such as the hook and filament, limits the spatiotemporal resolution. In this chapter, we describe a bead assay optimized for the analysis of the flagellar motor dynamics at near zero load. Key words Bacterial flagellar motor, Single-molecule measurement, Bead assay, Hook, Gold nanoparticle

1

Introduction The bacterial flagellar motor is a rotary nanomachine that rotates a long helical filament that extends in the cell exterior to propel bacteria in an aqueous milieu. The torque-generation mechanism of the flagellar motor is expected to be a design foundation for a highly efficient artificial nanomachine, but it remains unclear. Earlier studies investigated output properties of the flagellar motor by the tethered cell assay, where the rotation of each cell body attached to a glass surface via a flagellar filament is observed [1]. Since the cell body imposes a heavy load on the flagellar motor, the speed range of the tethered cell rotation is limited to ~10 Hz. In contrast, the bead assay performed by labeling a partially sheared flagellar filament with a microbead allows us to change loads in a bead-size-

Tohru Minamino et al. (eds.), Bacterial and Archaeal Motility, Methods in Molecular Biology, vol. 2646, https://doi.org/10.1007/978-1-0716-3060-0_11, © The Author(s), under exclusive license to Springer Science+Business Media, LLC, part of Springer Nature 2023

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Fig. 1 Rotation assay of the flagellar motor using a gold nanoparticle (φ60 nm). (a) A gold particle is attached to a sticky filament stub in a conventional bead assay. (b) The Salmonella mutant MMK2798iC, which lacks the flagellar filament and produces the elongated, straightened, and rigid hook. The electron micrograph on the right shows negatively stained hooks isolated from MMK2798iC. A gold nanoparticle is directly attached to the hook with three surface-exposed cys residues. OM, PG, and CM indicate the outer membrane, peptidoglycan layer, and cytoplasmic membrane, respectively

dependent manner (Fig. 1a) [2, 3]; e.g., the Salmonella flagellar filament labeled with 100 nm fluorescent beads rotates at ~250 Hz [4]. Labeling submicron-diameter beads enabled us to observe stepwise movements in the motors rotating at a few Hz, which reflect an elementary process of the mechanochemical energy coupling reaction [5, 6]. To analyze the motor dynamics at near zero load, a 60 nm-diameter gold nanoparticle was attached to the flagellar hook of E. coli using polyclonal anti-hook antibody, and the flagellar motor rotated at a maximum speed of ca. 300 Hz [7]. The bead assay is the most preferred method for single-molecule measurement of the bacterial flagellar motor over a wide range of external loads, but some concerning factors that reduce the measuring accuracy remain. Significant factors are attributed to the flexibility of the axial parts of the flagellum. First, a microbead attached to a partially sheared sticky flagellar filament (see Note 1) sometimes shows unstable rotational trajectories, although it depends on the position of bead attachment. Second, the probe movement could delay from the motor rotation with the response time τ = γ/κ, where

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γ is the drag coefficient and κ is the rigidity of the hook located between the motor and filament to act as a universal joint to transmit torque produced by the motor to the filament. To resolve these problems, we have constructed a Salmonella mutant strain, MMK2798iC, having a short, straight, rigid hook without filament by the following genetic engineering (Fig. 1b) [8]: 1. The fliC gene encoding the flagellin protein is knockout so that flagellar filament is not produced. 2. While the length of the native hook is regulated to be ~55 nm in Salmonella, the mutant hook is ~115 nm long due to an insertion in the FliK ruler protein, which measures the hook length during hook assembly [9]. Therefore, a 60 nm gold nanoparticle attached to the 115 nm hook does not interfere with flagellar motor rotation by its possible interaction with the cell surface. 3. The rod is a straight, stiff axial part and functions as a drive shaft of the flagellar motor [10]. The rod protein FlgG shows considerable sequence and structural similarities to the hook protein FlgE [11]. Eighteen amino-acid residues (YQTIRQPGAQSSEQTTLP) in FlgG, which are not in FlgE, confer the straightness and rigidity on the rod [11]. To make the hook straight and rigid, these 18 residues of FlgG are inserted between Phe-42 and Phe-43 of the hook protein FlgE [12]. 4. For direct binding of a gold nanoparticle to the hook, surface exposed three residues of FlgE, Thr-220, Thr-223, and Thr-224, are replaced by Cys residues. As a result, the MMK2798iC strain produces a rigid, straight, twice longer hook without the filament at the hook tip. This chapter describes the protocol to observe flagellar motor rotation at an extremely low load close to zero using the MMK2798iC strain.

2

Materials

2.1

Bacterial Strain

1. Salmonella enterica MMK2798iC [8].

2.2

Media

1. L-broth (LB): 10 g tryptone, 5 g yeast extract, 5 g NaCl per liter.

serovar

Typhimurium

strain

2. Motility medium: 10 mM potassium phosphate (pH 7.0), 0.1 M EDTA, 10 mM L-sodium lactate. 2.3 Cell Growth and Harvest

1. Shakers. 2. Test tubes. 3. Microcentrifuge. 4. 1.5 mL Eppendorf tubes.

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Probe

1. Gold nanoparticle (φ60 nm). 2. Sonicator. 3. 1.5 mL Eppendorf tubes.

2.5 Dark-Field Microscope

1. Upright microscope. 2. 100× oil immersion objective. 3. 5× relay lens. 4. Mercury fiber illumination system. 5. High-speed video camera (see Note 2).

2.6

Flow Chamber

1. Glass slide (0.8 ~ 1.0 mm in thickness). 2. 22 × 24 mm coverslip (borosilicate, noncoated glass, 0.13 ~ 0.17 mm in thickness). 3. Double-sided tape.

3

Methods

3.1 Preparation of a Flow Chamber

1. Soak coverslips in 10 N KOH for 5 h. 2. Wash coverslips with ultrapure water. 3. Store in 100% ethanol at room temperature. 4. Immediately before use, dry the washed coverslips. 5. Attach a glass slide with the washed coverslip with double-sided tape (Fig. 2).

3.2 Preparation of Gold Nanoparticles

1. Dilute gold nanoparticle solution into the motility medium (1: 10 dilution). 2. Ultrasonicate this solution to disperse aggregated gold nanoparticles (see Note 3).

Fig. 2 Schematic diagram of the flow chamber and optical setting for the bead assay. (a) Flow chamber. (b) Bacterial cells attached to the coverslip are observed through a high-speed video camera

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1. Culture the Salmonella MMK2798iC strain in LB at 37 °C overnight with shaking. 2. Dilute the overnight culture into fresh LB (1:100), and incubate at 37 °C for 3 h with shaking. 3. Transfer 1.0 mL of the bacterial culture into 1.5 mL Eppendorf tube, and centrifuge at 10,000g for 1 min at room temperature. 4. Suspend the cell pellet into a 250 μL volume of the motility medium. 5. Infuse the cell suspension into a flow chamber, and incubate for 20 min at room temperature. 6. Remove floating bacteria by running the motility medium into the flow chamber. 7. Infuse the suspension of gold nanoparticles into the flow chamber at room temperature (see Note 4). 8. Remove floating gold nanoparticles that did not attach to the hooks of MMK2798iC by running the motility medium.

3.4 Rotation Measurements

1. Place the flow chamber on a microscope stage. 2. Search gold nanoparticles rotating over the bacterial cell body, and record movie images at a frame rate of 5 kHz (see Note 2). 3. Analyze the recorded movies using a commercial or custombuilt software; Fig. 3 shows an example dataset analyzed by ImageJ and Microsoft Excel.

Fig. 3 Example data of bead rotation assay using MMK2798iC and φ60 nm gold nanoparticle. Tracking the bead position with ImageJ software (a) is followed by the analysis with Microsoft Excel (b). In (b), the X–Y trajectory in the top left panel shows a circular bead movement with a radius of ~50 nm, and the FFT spectrum (bottom left) of the sinusoidal curves showing the temporal change of the bead position in X (top right) or Y (bottom right) shows a broad peak at around 400 Hz, indicating the average rotation rate of the motor with a large fluctuation

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Notes 1. Salmonella flagellin, FliC, consists of four structural domains, D0, D1, D2, and D3. The flagellar filament lacking residues 205 through 293 of FliC, corresponding to the outer domain D3, shows a strong binding affinity for polystyrene beads and gold nanoparticles. Since this “sticky” filament realizes efficient probe labeling, the fliCΔ(205–293) allele is often used for conventional bead assays [13]. 2. The recording frame rate depends on the motor speed and the signal intensity (brightness) of gold nanoparticles. The MMK2798iC motor labeled with a 60 nm gold nanoparticle rotates at ~400 Hz, and it was recorded using a CMOS (complementary metal-oxide semiconductor) video camera at a frame rate of 5000 Hz [8]. 3. There is no strict standard for the condition of ultrasonication. We usually sonicate the bead solution for 10 min at 45 kHz. 4. Figure 4 depicts the way to infuse bead solution into a flow chamber. By tilting the flow chamber, the gravity-dependent slow passage of beads enables efficient bead labeling.

Fig. 4 Method for bead labeling. Infuse about 100 μL the bead solution in total, but divide it into two halves for two times infusion

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Acknowledgments We thank Yusuke V. Morimoto, Keiichi Namba, Nobunori Kamiike, and Koichi Hiraoka for their technical supports. This research has been supported by MEXT KAKENHI Grant Number JP25117501 to S.N. and JP20H05532 and JP22H04844 to T.M. and JSPS KAKENHI Grant number JP24770141 to S.N. and JP19H03182, JP22H02573, and JP22K19274 to T.M. References 1. Silverman M, Simon M (1974) Flagellar rotation and the mechanism of bacterial motility. Nature 249:73–74. https://doi.org/10. 1038/249073a0 2. Ryu WS, Berry RM, Berg HC (2000) Torquegenerating units of the flagellar motor of Escherichia coli have a high duty ratio. Nature 403:444–447. https://doi.org/10.1038/ 35000233 3. Chen X, Berg HC (2000) Torque-speed relationship of the flagellar rotary motor of Escherichia coli. Biophys J 78:1036–1041. https:// doi.org/10.1016/S0006-3495(00)76662-8 4. Nakamura S, Kami-ike N, Yokota JP et al (2009) Effect of intracellular pH on the torque-speed relationship of bacterial protondriven flagellar motor. J Mol Biol 386:332– 338. https://doi.org/10.1016/j.jmb.2008. 12.034 5. Sowa Y, Rowe AD, Leake MC et al (2005) Direct observation of steps in rotation of the bacterial flagellar motor. Nature 437:916–919. https://doi.org/10.1038/nature04003 6. Nakamura S, Kami-ike N, Yokota JP et al (2010) Evidence for symmetry in the elementary process of bidirectional torque generation by the bacterial flagellar motor. Proc Natl Acad Sci U S A 107:17616–17620. https://doi. org/10.1073/pnas.1007448107 7. Yuan J, Berg HC (2008) Resurrection of the flagellar rotary motor near zero load. Proc Natl Acad Sci U S A 105:1182–1185. https://doi. org/10.1073/pnas.0711539105

8. Nakamura S, Hanaizumi Y, Morimoto YV et al (2020) Direct observation of speed fluctuations of flagellar motor rotation at extremely low load close to zero. Mol Microbiol 113: 755–765. https://doi.org/10.1111/mmi. 14440 9. Minamino T (2018) Hierarchical protein export mechanism of the bacterial flagellar type III protein export apparatus. FEMS Microbiol Lett 365:fny117. https://doi.org/ 10.1093/femsle/fny117 10. Nakamura S, Minamino T (2019) Flagelladriven motility of bacteria. Biomolecules 9: 2 7 9 . h t t p s : // d o i . o r g / 1 0 . 3 3 9 0 / biom9070279 11. Fujii T, Kato T, Hiraoka KD et al (2017) Identical folds used for distinct mechanical functions of the bacterial flagellar rod and hook. Nat Commun 8:14276. https://doi.org/10. 1038/ncomms14276 12. Hiraoka KD, Morimoto YV, Inoue Y et al (2017) Straight and rigid flagellar hook made by insertion of the FlgG specific sequence into FlgE. Sci Rep 7:46723. https://doi.org/10. 1038/srep46723 13. Che Y-S, Nakamura S, Kojima S et al (2008) Suppressor analysis of the MotB(D33E) mutation to probe bacterial flagellar motor dynamics coupled with proton translocation. J Bacteriol 190:6660–6667. https://doi.org/10.1128/ JB.00503-08

Chapter 12 Live-Cell Fluorescence Imaging of Magnetosome Organelle for Magnetotaxis Motility Yukako Eguchi and Azuma Taoka Abstract The assessment of intracellular dynamics is crucial for understanding the function and formation process of bacterial organelle, just as it is for the inquisition of their eukaryotic counterparts. The methods for imaging magnetosome organelles in a magnetotactic bacterial cell using live-cell fluorescence imaging by highly inclined and laminated optical sheet (HILO) microscopy are presented in this chapter. Furthermore, we introduce methods for pH imaging in magnetosome lumen as an application of fluorescence magnetosome imaging. Key words Bacterial organelle, Magnetosome, Magneto-sensing, Magnetotactic bacteria, Magnetotaxis, Live-cell fluorescence imaging, pH imaging

1

Introduction Bacteria have several types of subcellular compartments, which have recently been called “bacterial organelles” (reviewed in refs. 1–3). Bacterial organelles, like eukaryotic organelles, have specific functions and distinct structures. Some bacterial organelles are bounded by a lipid membrane (e.g., magnetosomes, anammoxosomes, ferrosomes, and chromatophores), while others have proteinaceous boundaries (e.g., carboxysomes and metabolosomes) or a phase-defined membraneless boundary (e.g., nucleolus-like compartments). Although bacterial organelles are nanometer-sized structures in small bacterial cells, developments of advanced electron microscopic techniques, especially electron cryotomography, have enabled them to be discovered and their detailed ultrastructure to be explored [4–6]. Cryo-electron microscopy captures nearnative intracellular structures, but these are snapshot still-images of frozen cells in vitreous ice. Hence, live-cell fluorescence imaging is effective as a complementary technique for assessing intracellular

Tohru Minamino et al. (eds.), Bacterial and Archaeal Motility, Methods in Molecular Biology, vol. 2646, https://doi.org/10.1007/978-1-0716-3060-0_12, © The Author(s), under exclusive license to Springer Science+Business Media, LLC, part of Springer Nature 2023

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dynamics of organelles in cells throughout the cell cycle to gain a comprehensive understanding of bacterial organelles. Bacteria have various types of motilities [7], and some bacterial organelles are associated with them. Magnetosome is one of the conspicuous instances of the organelle that play a role in guiding the swimming direction of a bacterial cell. Magnetotactic bacteria synthesize magnetosomes to detect and swim along the geomagnetic field (reviewed in refs. 8–12). Magnetosome is a lipid bilayer membrane-enclosed organelle consisting of membrane vesicles derived from inner membrane invagination, a set of proteins, and magnetite (Fe3O4) or greigite (Fe3S4) crystals. Magnetosomes are aligned into a chain-like configuration localized along the central axis of cells and function as a cellular magnetic compass. Magnetotactic bacteria can orient themselves to find a favorable chemically stratified environment, such as a microaerobic habitat, due to the integration of magnetosomes in cellular motility. Due to the inclination of the Earth’s magnetic field, magnetotactic bacteria are able to fix their swimming direction into one dimension along the vertical axis in aquatic habitats. As a result, magnetotactic bacteria could effectively search for their favorable environment, which is vertically stratified, by combining magnetotaxis and aerotaxis/ chemotaxis [13]. Live-cell fluorescence imaging has been repeatedly used for investigating magnetosomes. For example, imaging of magnetosome dynamics has revealed the segregation process of magnetosome to daughter cells [14, 15]. Furthermore, fluorescence imaging of magnetosome-associated proteins has played an essential role in understanding how magnetosome-associated proteins organize magnetosomes (as examples of recent studies [16–18]). In this chapter, we introduce methods for fluorescence imaging of magnetosome dynamics using long-term time-lapse imaging. The method of pH imaging in the magnetosome lumen using a pH-sensitive fluorescence protein [19] is introduced as an application of live-cell fluorescence imaging. In this study, we have used the α-proteobacterial magnetotactic bacterium Magnetospirillum magneticum AMB-1 [20] (AMB-1) as a specimen. The AMB-1 strain is one of the model species of magnetotactic bacteria and allows for detailed genetic analysis [21].

2

Materials

2.1 Preparation for AMB-1 Cells Expressing GFP-Fused Magnetosome Membrane Proteins

1. M. magneticum AMB-1 (ATCC 700264). 2. Escherichia coli strain S17-1 [22] or WM3064 [23] as a host strain of biparental conjugation. 3. A protein expression vector pBBR111 [24] for M. magneticum AMB-1.

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4. Magnetospirillum growth medium (MSGM) [23] containing per liter: 5 mL of Wolfe’s mineral solution (without FeSO4), 0.68 g of KH2PO4, 0.12 g of NaNO3, 0.07 g of sodium acetate, 0.035 g of ascorbic acid, 0.37 g of tartaric acid, 0.37 g of succinic acid, and 0.05 g of sodium thiosulfate. Adjust pH to 6.9 with NaOH, and autoclave. Before inoculation, add 1% (v/v) of Wolfe’s vitamin solution and 1% (v/v) of 3 mM ferric malate to medium. For agar plates, add 7 g of agar per liter of MSGM. 5. Wolfe’s mineral solution containing per liter: 1.5 g of nitrilotriacetic acid, 3.0 g of MgSO4 7H2O, 0.5 g of MnSO4, 1.0 g of NaCl, 0.1 g of CoCl2 6H2O, 0.1 g of CaCl2, 0.1 g of ZnSO4 7H2O, 0.01 g of CuSO4 5H2O, 0.01 g of AlK(SO4)2 12H2O, 0.01 g of H3BO3 0.01 g, 0.01 g of Na2MoO4 2H2O, and 4.88 mg of Na2SiO3. Firstly, dissolve nitrilotriacetic acid in approximately 500 mL of water, adjust pH to about 6.5 with KOH, and then add the following compounds in the order described above. Autoclave the solution, and store it at 4
C until use. 6. Wolfe’s vitamin solution containing per liter: 2.0 mg of biotin, 2.0 mg of folic acid, 10.0 mg of pyridoxine HCl, 5.0 mg of thiamine HCl, 5.0 mg of riboflavin, 5.0 mg of nicotinic acid, 5.0 mg of calcium D-(+)-pantothenate, 0.1 mg of cyanocobalamin, 5.0 mg p-aminobenzoic acid, and 5.0 mg of thioctic acid. Sterilize the solution using a 0.2 μm pore-sized filter, and store it at 4
C until use. 7. Ferric malate solution containing per liter: 0.486 g of FeCl3 and 1.21 g of DL-malic acid. Sterilize the solution using a 0.2 μm pore-sized filter, and store it at room temperature. 8. LB broth, for Escherichia coli cultivation, containing per liter: 10 g of Bacto tryptone, 5 g of Bacto yeast extract, and 10 g of NaCl. Adjust pH to 7.5 with 5 N NaOH, and sterilize by autoclave [25]. 9. Antibiotics are added to MSGM and LB broth at the following concentration: for AMB-1, kanamycin (5 μg/mL); for E. coli, kanamycin (20 μg/mL) and trimethoprim (100 μg/mL). 10. 60 mM diaminopimelate (DAP) solution sterilized by filtration. 11. Membrane filter, 0.45 μm pore size, mixed cellulose ester: cut a membrane filter to about 2 cm2, and autoclave them in advance for use. 12. Anaerobic jar (e.g., BBL GasPak BD, USA). 13. Gas pack for microaerobic atmosphere (e.g., Anaero Pack “MicroAero,” Mitsubishi Gas Chemical, Japan).

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2.2 Live-Cell Fluorescence Imaging of Magnetosome Positioning

1. Attofluor Cell Chamber for microscopy. 2. Round coverslip, 25 mm diameter, 0.12–0.17 mm thick: wash coverslips twice in 0.2 N NaOH for 10 min in ultrasonication bath. Then, rinse them two times with 99.5% (v/v) EtOH, and store them in 99.5% (v/v) EtOH until use. 3. Gellan gum gel pads: Add 0.27 g gellan gum powder and 50 mL MSGM in a 100 mL flask. Then, dissolve gellan gum using a microwave. After that, add 0.5 mL of Wolfe’s vitamin solution, 0.5 mL of ferric malate solution, 5 μg/mL (final conc.) kanamycin, and 25 μL of 2 M MgCl2 to solidify gellan gum. Finally, pour 3.5 mL of the resulting gellan gum solution into a small square-shaped weighting tray (4  4 cm) and solidify at room temperature. 4. A total internal reflection fluorescence (TIRF) microscopybased system with an inverted microscope (e.g., Ti-E, Nikon, Japan) equipped with a 100  TIRF objective (e.g., Nikon, Japan), a 1.5 C-mount adapter (e.g., Nikon, Japan), and a 488 nm laser (e.g., Sapphire, Coherent, USA). 5. A high-sensitivity electron-multiplying charge-coupled device (EMCCD) camera (e.g., iXon3; Andor, UK). 6. Stage incubator for microscope (e.g., Toukai Hits, Japan). 7. Microscope operating software (e.g., NIS Elements AR software Nikon, Japan).

2.3 Live-Cell pH Measurements in Magnetosome Lumen

1. Calibration buffers with desired pH ranging from 5 to 9: Mix 0.1 M citric acid and 0.2 M Na2HPO4 in proper mixing ratios [26]. Add 20 mM sodium benzoate (final conc.) to each mixture. Before use, measure the actual pH of calibration buffers at 25
C using a pH meter. 2. A grating spectrometer (e.g., CLP-50; Andor, UK): set between an inverted microscope (e.g., Ti-E, Nikon, Japan) and a high-sensitivity EMCCD camera (e.g., iXon3; Andor, UK). 3. A filter block equipped with a dichroic filter (e.g., Di02-R48825x36; Semrock) and a long-pass emission filter (e.g., BLP01488R-25; Semrock). 4. Round coverslip, 18 mm diameter, 0.12–0.17 mm thick (e.g., Matsunami, Japan). 5. Shading hood for the inverted microscopic stage.

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Methods

3.1 Preparation for AMB-1 Cells Expressing GFP-Fused Magnetosome Membrane Proteins

1. Green fluorescence protein (GFP) fusions to magnetosome membrane protein are used for fluorescence labeling of magnetosomes [27]. A fusion gene of a magnetosome membrane protein (e.g., mamC, mmsF, and mamI) with gfp is cloned into the conjugative broad-host-range protein expression vector pBBR111 [24] harboring the tac promoter (Fig. 1) (see Notes 1 and 2). 2. Prepare a conjugative donor E. coli strain S17-1 [22] or WM3064 [23], containing the plasmid prepared in step 1.

Fig. 1 Fluorescence labeling of magnetosomes. (a) Examples of magnetosome membrane protein used as magnetosome marker for fluorescence imaging. MamC regulates the size and shape of magnetite crystals in magnetosomes, while MamI is essential for the formation of magnetosome membrane vesicles. The mineralizing protein MamC can be used to specifically detect the positions of mineral-containing magnetosomes, whereas MamI can be used to detect magnetosome vesicles with and without magnetite. (b) Expression vector for GFP-fused magnetosome membrane protein is transferred to M. magneticum AMB-1 by biparental conjugation

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3. Cultivate a recipient strain, AMB-1 or its derived strain of interest, in 400 mL of MSGM for 3 days at 28
C in a 500 mL glass medium bottle. After inculcation, exchange the headspace of the bottle to 1% (v/v) O2—99% (v/v) N2 gas to keep microaerobic conditions suitable for AMB-1 growing. Cultivate until the culture reaches the early stationary phase (see Note 3). 4. Cultivate a conjugative donor E. coli strain S17-1 or WM3064, containing the plasmid for GFP-fused magnetosome membrane protein in 4 mL of LB medium containing kanamycin at 37
C for overnight (see Note 4). 5. Centrifuge 400 mL of the recipient AMB-1 culture at 8000  g for 10 min at 25
C (see Note 5). Suspend the pellet with about 4 mL MSGM. 6. Centrifuge 4 mL of the donor E. coli strain culture at 8000  g for 5 min at 25
C. 7. Suspend the E. coli pellet in 10 mL of MSGM (containing 0.3 mM DAP for WM3064). Then, centrifuge the suspension again at 8000  g for 10 min at 25
C, and suspend the pellet in 10 mL MSGM. Repeat this process twice to wash out the antibiotics from E. coli cells. Finally, suspend the E. coli pellet in 2 mL of MSGM (containing 0.3 mM DAP for WM3064). 8. Add 0.5 mL of the AMB-1 cell suspension and 0.5 mL of the E. coli cell suspension in a 1.5 mL tube, and centrifuge the mixture for 5 min at 25
C. Suspend the resulting pellet with 50 μL MSGM. 9. Put a piece of a membrane filter (pore size: 0.45 μm) on the MSGM agar plate (containing 0.3 mM DAP for WM3064), and then put 50 μL of the resulted suspension on the nitrocellulose filter. The donor and recipient cells are highly concentrated and stick together on the membrane filter. 10. Put the agar plate in a jar with a microaerobic atmosphere (e.g., using BD BBL GasPak jar containing Anaero Pack “MicroAero”), and incubate for 6 h at 28
C. 11. Suspend cells on the filter with 100 μL of MSGM, and transfer to a 1.5 mL tube. Prepare serial dilutions of cell suspensions, and inoculate the dilutions into kanamycin-containing MSGM agar plates. Incubate the agar plates in the jar with the microaerobic atmosphere at 28
C for about 7 days to grow AMB-1 colonies. 12. Take some of the colonies and cultivate in 1.5 mL of MSGM with kanamycin, and use the cultivated cells for fluorescence imaging as described in Subheading 3.2.

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3.2 Live-Cell Imaging of Magnetosome Positioning

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1. Soak 25 mm coverslip in 0.005% (v/w) poly-L-lysine solution, and dry up the coverslip in air. Set the coverslip to an Attofluor Cell Chamber. 2. Cultivate the AMB-1 derived strain expressing GFP-fused magnetosome membrane protein in MSGM with kanamycin for 20–40 h until the late logarithmic growth phase. 3. Place the gellan gum gel and 10 mL of fresh MSGM containing Wolf’s vitamin solution, ferric malate solution, and kanamycin in a microaerobic atmosphere overnight to equilibrate (see Note 6). 4. Figure 2 depicts a schematic of the sample setup for magnetosome imaging using a highly inclined and laminated optical sheet (HILO) microscope. At first, add about 500 μL of the culture on the poly-L-lysine-treated coverslip in the Attofluor Cell Chamber. Carefully place a 15  15 mm2, 5 mm thick gellan gum gel pad on the coverslip to sandwich the cells between the bottom coverslip and the gellan gum gel pad. This prevents cell movement during long-term time-lapse imaging (see Note 7). Remove the excess culture from the

Fig. 2 Schematics of the sample setup for live-cell imaging of magnetosomes in M. magneticum AMB-1 cells. Logarithmic growth cells expressing the magnetosome membrane protein and GFP fusion protein are immobilized on a poly-Llysine set-coated coverslip in an Attofluor cell chamber. To prevent cells from moving or detaching during observation, a pad of gellan gum gel is placed on the cells. The inside of the chamber is filled with MSGM, and another coverslip is used to cover the chamber. After that, time-lapse observation using oblique illumination in a HILO microscope is performed to image magnetosome localization

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edge of the chamber using a pipette. By this, cells are well sandwiched between the gel and the coverslip. Then, add fresh MSGM to fill up the chamber. Put another 25 mm coverslip on the top of the chamber as a cover. No air bubbles should be inside the chamber. Using a pipette tip, tightly attach the upper coverslip to the chamber, and remove excess medium that has overflowed on the coverslip. Then, to prevent drying and maintain microaerobic conditions, apply a fast-drying topcoat along the side of the upper coverslip. 5. Set the sample to a stage incubator equipped with a TIRF microscopy-based inverted microscope. The microscope is set to operate in highly inclined and laminated optical sheet (HILO) microscopic mode. 6. Maintain the sample temperature in the stage incubator at 28
C. Before starting time-lapse imaging, cultivate cells in the chamber for 4–6 h to grow AMB-1 cells in advance. Using bright-field observation, check for cell divisions occurring in the observing area (see Note 8). 7. HILO imaging setup: a 488 nm laser is used to illuminate the sample at an oblique angle that is slightly steeper than the critical angle required for total reflection to illuminate entire bacterial cells. The laser beam angle is adjusted manually to optimize the signal-to-noise ratio. 8. Take a series of time-lapse images using the operation software of the microscope (e.g., NIS Elements AR software, Nikon, Japan) (Fig. 3). Typical exposure times for GFP and brightfield images are 300 ms and 100 ms, respectively. The interval of time-lapse is 1 min. The samples are illuminated only during exposure. The focus is kept using a focus maintaining system equipped with the microscope (e.g., Perfect Focus System equipped with a Ti-E microscope, Nikon, Japan). 3.3 Live-Cell pH Measurements in Magnetosome Lumen

1. Prepare a pBBR111-derived plasmid for expressing a pH-sensitive fluorescent protein, E2GFP [28], fused with the C-terminal of a magnetosome membrane protein, Mms6 (see Notes 9 and 10). Then, transfer the plasmid to AMB-1 as described in Subheading 3.1. 2. To prepare the pH standards cells, add aliquots (1 mL) of AMB-1 culture expressing E2GFP fusion into 1.5 mL tubes, and centrifuge at 8000  g for 5 min at 25
C. Then, suspend the pellets with 0.5 mL of each calibration buffer ranging from pH 5 to 9. Repeat these steps two times to wash out culture media. 3. To prepare the specimen cells for pH measurement, add 1 mL of AMB-1 derived cells of interest expressing E2GFP fusion

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Fig. 3 Examples of long-term time-lapse imaging of magnetosomes. MamC-GFP is used as a magnetosome marker. (Top) Magnetosomes in wild-type cells show patchy spots localized along the long axis of the cell. The magnetosome localization is maintained stably during the entire cell cycle as indicated by the parallel lines in the kymographs of GFP fluorescence. (Bottom) In contrast to the static straight-chain observed in wild-type cells, the magnetosomes in a deletion mutant of the magnetosomal cytoskeleton protein MamK randomly move throughout the cell, forming small, fast-moving fluorescence foci or large slow-moving fluorescence foci

into a 1.5 mL tube, and centrifuge at 8000  g for 5 min at 25
C. Suspend the pellet with 0.5 mL of fresh MSGM, and repeat these steps two times. 4. To prepare the sample for imaging, place 20 μL of the cell suspension on a poly-L-lysine-coated coverslip in an Attofluor Cell Chamber. Cover the sample with an 18 mm coverslip, then close the Attofluor Cell Chamber with a 25 mm coverslip, and apply a fast-drying topcoat along the side of the upper coverslip to prevent sample drying. 5. Set a grating spectrometer between the inverted TIRF microscope and the EMCCD camera (Fig. 4). To detect the fluorescence spectrum, set a long-pass emission filter to a filter block. 6. Set the chamber containing sample cells to the stage. Adjust the microscopic setup as described in step 7 of Subheading 3.2. Adjust the position of the stage to allow a target cell to be placed at the center of the microscope field. Then, insert the 0.1 mm slit of the grating spectrometer (Fig. 4). If the target cells are out of the slit, readjust the position of the stage. 7. Cover the stage with the shading hood. Insert a grating of the spectrometer, and take an emission fluorescence spectral image at wavelengths from 500 to 600 nm by exposing for 20 s at

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Fig. 4 Microscopic fluorescence spectroscopy. (a) The grating spectrometer equipped with a TIRF-based microscope. (b) Schematics of obtaining the fluorescence spectrum from a single cell. (i) Fluorescence image of a target cell (white arrowhead) in the slit of the grating spectrometer. (ii) An emission fluorescence spectral image obtained from the setup shown in panel (i). The x-axis of the image represents emission wavelength. The y-axis of the image is the same as in panel (i). The spectrum from the emission fluorescence signal of the target cell is shown in a blue dashed rectangle. (iii) The intensity line profile of the blue dashed rectangle in panel (ii). This profile is used as the emission spectrum of the target cell

488 nm excitation. The intensity line profile from the emission fluorescence spectral image represents the emission spectrum from the cell (Fig. 4b). Obtain emission spectra for at least three cells per condition (see Note 11). 8. Repeat steps 4–8 using each pH standard cells and the specimen cells of interests as samples. 9. Calculate the R488 nm values of each spectrum from the pH standards cells and the specimen cells. R488 nm is the ratio of the sum of fluorescence intensities ranging from 522 to 524 nm to those ranging from 508 to 510 nm (Fig. 5). 10. Plot the R488 nm values obtained from each pH standards cell versus the pH value of the calibration buffers. Fit the obtained plot by a least-squares fitting with the following Eq. 1.

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Fig. 5 Schematics of emission spectra obtained from E2GFP expressing AMB-1 cells in a pH range of 5–9. The ratio of the sum of fluorescence intensities at wavelengths ranging from 522 to 524 nm (blue arrow) to those at wavelengths ranging from 508 to 510 nm (red arrow) is used to estimate the pH of the magnetosome lumen



R488 nm

0

Rf þ 10ðpK pH Þ ¼ R0  0 1 þ 10ðpK pH Þ

 ð1Þ

Determine the parameters of R0, Rf, and pK0 by fitting. 11. Plug in the R488 nm value obtained from the specimen cells for the following Eq. 2 to calculate the pH value.

  R0  Rf  R488 nm pH ¼ pK  log 10 R488 nm  R0 0

4

ð2Þ

Notes 1. Magnetosomes consist of multiple types of specifically localized membrane-integral proteins [29]. The magnetosome membrane proteins, MamC [15, 27, 30], MmsF [18], and MamI [15, 30], are used as magnetosome markers. Magnetite biomineralization associating proteins, such as MamC and MmsF, can be used to image magnetite-containing matured magnetosome, while magnetosome vesicle formation process associating proteins, such as MamI, seem to be capable of imaging the entire magnetosome localization, including immature and matured ones. 2. The broad-host-range plasmid derived from pBBR1 [31] is available as a replicative plasmid for expressing recombinant proteins in AMB-1. The plasmid can be transferred from the conjugative E. coli host strain by biparental conjugation.

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3. Absorbance at 600 nm of an early stationary culture of AMB-1 wild-type is about 0.1. 4. WM3064 can grow in the presence of DAP. Add 0.3 mM DAP (final conc.) in LB broth for the cultivation of WM3064. 5. The cultures of AMB-1 and E. coli should not be cooled during centrifuge to maintain their cell activity. 6. Manipulations of the steps 3 and 4 in the Subheading 3.2 should be performed in a microaerobic atmosphere (1–10% (v/v) oxygen), such as a globe box or under flowing 1–10% (v/v) oxygen-containing nitrogen-based gas. 7. Agar pad might interfere the focus maintaining system (e.g., Perfect Focus System, Nikon, Japan) due to turbidity of solidified agar. Gellan gum gel is fully transparent and more suitable for maintaining focus during long-term observation. 8. Cell divisions are examined to confirm the healthiness of observed cells. Healthy cells can be selected by this observation. 9. The pH-sensitive fluorescence protein E2GFP demonstrates spectral forms that are convertible upon pH changes during both excitation and emission, with the pK values close to 7.0 [28]. 10. Mms6, a magnetosome membrane protein, is used to direct E2GFP into the magnetosome lumen. Because the C-terminal region of Mms6 is involved in magnetite crystal shape regulation [32, 33], C-terminally fused E2GFP is positioned in the magnetosome lumen. 11. The spectrum obtained from the area where no cell is found should be used as the background. Subtract the background spectrum from the spectrum obtained from a cell to eliminate the effects of filters.

Acknowledgments We thank Dr. A. Komeili (University of California, Berkeley) for providing the mamK deletion mutant of M. magneticum AMB-1 and invaluable advice. We also appreciate Dr. L.F. Wu (Aix-Marseille Universite´) providing plasmid pBBR111. This work was supported by a MEXT KAKENHI Grant Number 24117007, and JSPS KAKENHI 21KK0126, 20K15430, 19H02868, and 17KK0145.

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31. Antoine R, Locht C (1992) Isolation and molecular characterization of a novel broadhost-range plasmid from Bordetella bronchiseptica with sequence similarities to plasmids from gram-positive organisms. Mol Microbiol 6(13):1785–1799 32. Amemiya Y, Arakaki A, Staniland SS et al (2007) Controlled formation of magnetite crystal by partial oxidation of ferrous hydroxide in the presence of recombinant magnetotactic bacterial protein Mms6. Biomaterials 28(35): 5381–5389 33. Arakaki A, Webb J, Matsunaga T (2003) A novel protein tightly bound to bacterial magnetic particles in Magnetospirillum magneticum strain AMB-1. J Biol Chem 278(10): 8745–8750

Chapter 13 Swarming Motility Assays in Salmonella Jonathan D. Partridge and Rasika M. Harshey Abstract Salmonella enterica has six subspecies, of which the subspecies enterica is the most important for human health. The dispersal and infectivity of this species are dependent upon flagella-driven motility. Two kinds of flagella-mediated movements have been described—swimming individually in bulk liquid and swarming collectively over a surface substrate. During swarming, the bacteria acquire a distinct physiology, the most significant consequence of which is acquisition of adaptive resistance to antibiotics. Described here are protocols to cultivate, verify, and study swimming and swarming motility in S. enterica, and an additional “border-crossing” assay, where cells “primed” to swarm are presented with an environmental challenge such as antibiotics to assess their propensity to handle the challenge. Key words Flagella, Motility, Surface motility, Swarming, Swimming

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Introduction A diverse array of bacteria depend on flagella-mediated motility to move through their environment and reach appropriate host targets for infection [1–7]. The flagellum is an external organelle that translates a rotary force into propulsion where the force is generated by ion gradients across the cell membrane [4, 8–12]. In the peritrichously flagellated and well-studied bacteria Escherichia coli and S. enterica, the flagellar machinery is coupled to a chemotaxis sensory apparatus, whose output modulates the direction of the rotation of the motor to which the flagellum is anchored in the inner membrane [13–15]. Bidirectional switching of the motor causes flagella bundling and unbundling, resulting in forward “runs” and cell-reorienting “tumbles” respectively, promoting bacterial movement toward positive stimuli such as nutrients, or away from negative stimuli such as toxic substances [16–18]. Movement of individual bacteria through bulk liquid is called swimming, where chemotaxis plays a major role in fostering bacterial colonization of optimal environments [8, 18–20]. Swarming, the focus of this article, employs the same flagellar machinery as during

Tohru Minamino et al. (eds.), Bacterial and Archaeal Motility, Methods in Molecular Biology, vol. 2646, https://doi.org/10.1007/978-1-0716-3060-0_13, © The Author(s), under exclusive license to Springer Science+Business Media, LLC, part of Springer Nature 2023

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swimming, but here the bacteria move on a surface, and not as individuals but as a collective [2, 21–27]. Chemotaxis is not required for swarming, but bacteria at the edge apparently do show chemotactic behavior [28]. Within the swarm, the bacteria have a different physiology; they exhibit faster speeds, an altered run-tumble bias that suppresses chemotaxis, and tolerance to levels of antibiotics that are lethal to swimmers [29–32]. Swarmers can be broadly grouped into two categories: temperate swarmers, which can swarm on “softer” media solidified with 0.5% (w/v) to 0.8% (w/v) agar, and robust swarmers, which can migrate across “hard” media solidified with >1% (w/v) agar [21, 27, 33]. The robust swarmers include Proteus mirabilis and Vibrio parahaemolyticus; these bacteria display multiple adaptations to enable movement across “hard” surface environments (secretion of surfactants and polysaccharides, cell elongation, hyperflagellation), where lower hydration and higher surface friction are expected [21, 27, 33]. Temperate swarmers include E. coli, Bacillus subtilis, S. enterica, Serratia. marcescens, and Pseudomonas aeruginosa; these bacteria often have either none or only a subset of the adaptations seen in robust swarmers and may require in addition sugars or special agar to support swarming [27]. General swarming assay protocols [34], as well as specifically tailored instructions for E. coli [35], B. subtilis [36], P. mirabilis [37], P. aeruginosa [38], and V. parahaemolyticus [39], among others, have been published. Many techniques developed for E. coli are frequently applied (successfully) to Salmonella due to the close evolutionary relationship of the two bacteria [40]. There are, however, notable differences in flagellar motility between these two species. Salmonella flagellar motors rotate at slower speeds and show fewer motor reversals than E. coli [41]. While both bacteria require inclusion of glucose in the swarm media, Salmonella are less fastidious in that they can swarm on Bacto agar while E. coli requires a special Eiken agar sold in Japan [42]. Salmonella colonize a surface faster than E. coli at either 30 or 37 °C, while E. coli are best assayed at 30 °C. Salmonella will swarm on media solidified with 0.4–0.7% (w/v) agar, while for E. coli, this range is slightly narrower with 0.4–0.5% (w/v) agar. Here, we describe three protocols for studying flagellamediated motility in Salmonella. For each, proper preparation of media and appropriate timing for drying are crucial to ensure consistency and proper establishment of bacterial swarms. Surface hydration is affected by agar type, agar thickness, drying temperature, incubation temperature, and overall humidity. The first is the well-established swimming (or chemotaxis) assay, where Salmonella inoculated at the center of a petri dish, containing rich media solidified with 0.3% (w/v) agar (“soft agar”) consume local nutrients to create a gradient, then climb the gradient via chemotaxis, and move outward; this assay measures both motility and

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chemotactic ability [19, 43]. The second is the swarm assay, where Salmonella swarm on the surface of media solidified with ~0.6% (w/v) agar. Finally, the border-crossing assay, where dualcompartment petri dishes are used to pour different media in the two compartments, connected at their border by an agar bridge. Typically, only one compartment includes a stressor such as antibiotics or oxidative stress. Swarming is initiated in the no-antibiotic compartment, and the ability of the swarm to cross the agar bridge and overcome the antibiotic challenge in the second compartment is assessed [27, 31, 35].

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Materials Ensure that all materials and reagents used in the experiments below are appropriately sterilized, either by filtration or autoclaving.

2.1 Swimming Motility Assay

1. Salmonella strain(s) of interest (see Note 1). 2. Lennox broth (LB): per liter ddH20, 10 g of tryptone, 5 g of yeast extract, and 5 g of NaCl (see Note 2). Autoclave to sterilize, with an exposure time of 20 min, and temperature of 121 °C. 3. Soft agar (chemotaxis assay): per liter, 10 g of tryptone, 5 g of yeast extract, 5 g of NaCl, and 3 g of agar (see Notes 2 and 3). Autoclave to sterilize, with an exposure time of 20 min, and temperature of 121 °C. Cool to 55 °C (see Note 4) before transferring 20 mL of agar to each petri dish. Use once fully cooled and set. Prepare soft agar plates for use on the day of the experiment (see Note 5). 4. Sterile culture tubes and caps. 5. Incubator (30 °C) with an associated shaker to grow bacterial cultures. 6. 100 mm × 15 mm sterile petri dishes. 7. Incubator (30 °C) with shelving to incubate swim plates. 8. Inoculating loop. 9. Pipettes and sterile tips for inoculating cultures/plates. 10. 2 L flask. 11. Magnetic hotplate stirrer and stir bar. 12. Spectrophotometer and 1.5 mL semi-micro cuvettes. 13. Stopwatch or timer. 14. Ruler. 15. System to capture images of motility plates (see Note 6).

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2.2 Swarming Motility Assay

1. Salmonella strain(s) of interest (see Note 1). 2. Lennox broth (LB): per liter ddH20, 10 g of tryptone, 5 g of yeast extract, and 5 g of NaCl (see Note 2). Autoclave to sterilize, with an exposure time of 20 min, and temperature of 121 °C. 3. Swarm agar: per liter, 10 g of tryptone, 5 g of yeast extract, 5 g of NaCl, and 6 g of Eiken agar (see Note 7). Autoclave to sterilize, with an exposure time of 20 min, and temperature of 121 °C. Cool to 55 °C (see Note 4), add glucose to a final concentration of 0.5% (w/v) (see Note 7), before transferring 20 mL of agar to each petri dish. Prepare swarm agar plates the day prior to use, and allow to dry at room temperature with lids in place overnight, or ~16 h (see Note 8). 4. Sterile culture tubes and caps. 5. Incubator (30 °C) with an associated shaker to grow bacterial cultures. 6. 100 mm × 15 mm sterile petri dishes. 7. Incubator (30 °C) with shelving to incubate swarm plates. 8. Inoculating loop. 9. Pipettes and sterile tips for inoculating cultures/plates. 10. 2 L flask. 11. Magnetic hotplate stirrer and stir bar. 12. Spectrophotometer and 1.5 mL semi-micro cuvettes. 13. Stopwatch or timer. 14. Ruler. 15. System to capture images of motility plates (see Note 6). 16. Microscopy system to visualize and verify swarm colony (see Note 9).

2.3 The BorderCrossing Swarming Motility Assay

1. Salmonella strain(s) of interest (see Note 1). 2. Lennox broth (LB): per liter ddH20, 10 g of tryptone, 5 g of yeast extract, and 5 g of NaCl (see Note 2). Autoclave to sterilize, with an exposure time of 20 min, and temperature of 121 °C. 3. Swarm agar: per liter, 10 g of tryptone, 5 g of yeast extract, 5 g of NaCl, and 6 g of Eiken agar (see Note 7). Autoclave to sterilize, with an exposure time of 20 min, and temperature of 121 °C. Cool to 55 °C (see Note 4), and add glucose to a final concentration of 0.5% (w/v) (see Note 7), before using to set up a dual-compartment petri dish (see Subheading 3.3 for details). Prepare border-crossing swarm agar plates the day prior to use, and allow to dry at room temperature with lids in place overnight, or ~16 h (see Note 8).

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4. Sterile culture tubes and caps. 5. Incubator (30 °C) with an associated shaker to grow bacterial cultures. 6. 100 mm × 15 mm sterile dual-compartment petri dishes. 7. Incubator (30 °C) with shelving to incubate swarm plates. 8. Inoculating loop. 9. A chosen agar supplement to challenge the swarming bacteria, e.g., antibiotics. 10. Pipettes and sterile tips for inoculating cultures/plates. 11. 2 L flask. 12. Magnetic hotplate stirrer and stir bar. 13. Spectrophotometer and 1.5 mL semi-micro cuvettes. 14. Stopwatch or timer. 15. Ruler. 16. System to capture images of motility plates (see Note 6).

3

Methods

3.1 Swimming Motility Assay

1. On the day prior to the assay, inoculate an overnight culture of Salmonella in 5 mL of LB, at 30 °C, with aeration. 2. On the day of the experiment, prepare soft agar plates. 3. Subculture 50 μL of Salmonella into 5 mL of LB (1:100 dilution), and grow at 30 °C, with aeration, to an OD600 value of ~0.6. 4. Gently pipette 6 μL of the culture into the center of a soft agar plate, the pipette tip penetrating the agar slightly, taking care to avoid contact with the bottom of the dish. Carefully withdraw the pipette back without disturbing the agar. 5. Carefully move the inoculated plates to a 30 °C incubator (see Note 10). Keep plates level. Do not invert (see Note 11), and do not stack plates on top of each other (see Note 12). 6. Monitor the plates periodically if necessary. Migration will be observable around the inoculation site after several hours (~3 h). Over time the bacteria will move outward, with the characteristic chemotactic rings becoming evident, eventually reaching the edges of the dish (~8 h) [44]. The time required for incubation will be dependent on the strain of interest. 7. Record the diameter with a ruler and the associated time taken to establish the ring (see Note 13). 8. Photograph the plates (see Note 6, and Fig. 1a for an example).

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Fig. 1 Observing swimming and swarming bacteria. (a) Salmonella swimming motility on a soft agar (0.3% [w/v] agar) plate, after 8 h incubation at 30 °C. (b) Salmonella swarming motility on a swarm agar (0.6% [w/v] agar, 0.5% [w/v] glucose) plate, after 6 h incubation at 30 °C. (c) Salmonella swarming motility across a border-crossing plate (0.6% w/v agar, 0.5% w/v glucose), after 8 h incubation at 30 °C. In this example, kanamycin (20 μg/mL) is added to the rightside compartment. (d) Visualization of an expanding Salmonella swarm front propagated as in (a), imaged from above using an Olympus microscope with a 40× objective

3.2 Swarming Motility Assay

1. On the day prior to the assay, inoculate an overnight culture of Salmonella in 5 mL of LB, at 30 °C, with aeration. 2. Prepare swarm-agar plates, and allow to dry overnight without stacking (see Note 12). 3. On the day of the experiment, subculture 50 μL of Salmonella into 5 mL of LB (1:100 dilution), and grow at 30 °C, with aeration, to an OD600 value of ~0.6. 4. Gently spot 6 μL of the culture atop and in the center of a swarm agar plate. Unlike in the instruction for swimming, avoid penetrating or disturbing the agar surface (see Note 14). 5. Allow culture to dry (~10 min) until liquid is absorbed into agar (see Note 8).

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6. Carefully move the inoculated plates to a 30 °C incubator (see Notes 10 and 15). Keep plates level, do not invert during incubation (see Note 11), and do not stack plates on top of each other (see Note 12). 7. Monitor the plates (see Note 16). After several hours (~3 h), migration will be observable around the inoculation site. Over time, the bacteria will move outward across the entire agar surface (see Note 17). The time required for incubation will depend on the strain of interest as well as the established plate/ environmental conditions (see Note 15). 8. Record the colony diameter at the widest point with a ruler and the time taken to establish the colony (see Note 18). 9. Image the plates (see Note 6, and Fig. 1b for an example). 10. If required, directly observe the swarming bacteria using a 10× objective or a long-working distance 40× objective to verify the characteristic whirls and flows associated with this type of motility (see Note 9). 3.3 The BorderCrossing Swarming Motility Assay

1. On the day prior to the assay, inoculate an overnight culture of Salmonella in 5 mL of LB, at 30 °C, with aeration. 2. Prepare border-crossing swarm-agar plates (see Note 19, and Fig. 2) by pouring ~30 mL of swarm agar with desired supplementation, in this instance, an antibiotic into the right-hand section of a dual-compartment petri dish. Ensure agar is level with the plastic divide between compartments but not overflowing between the compartments. Once the agar has hardened, fill the left-hand compartment with ~30 mL of regular swarm agar, again to the point of contact with the plastic divide. Before it sets, use a sterile pipette tip to gently drag the agar over the border, thereby connecting the two sides with

Fig. 2 Border-crossing swarm plates. A schematic showing the various stages involved in preparing for Salmonella border-crossing assays. From left to right, take a dual-compartment petri dish, pour ~30 mL of 0.6% [w/v] swarm agar (with 0.5% [w/v] glucose, and any desired challenge supplementation, e.g., antibiotic) to the right chamber until the chamber is level with the divider, and allow to set. Fill the left chamber with the same swarm agar, omitting the supplementation, until flush with the divider. Use a sterile pipette tip to gently drag the molten agar across the divider, thereby connecting the two sides with a ~1 mm tall bridge. Allow the completed plate to dry at room temperature overnight for use in the following day. For a representative image of Salmonella colonizing a border-crossing plate, see Fig. 1d

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a ~1 mm tall agar bridge (see Note 20). Allow to dry overnight without stacking (see Note 12). For a representative image of Salmonella colonizing a border-crossing plate, see Fig. 1d. 3. On the day of the experiment, subculture 50 μL of Salmonella into 5 mL of LB (1:100 dilution), and grow at 30 °C with aeration to an OD600 value of ~0.6. 4. Gently spot 6 μL of the culture on the surface of the agar and in the center of the left-hand compartment with normal swarm media. Avoid penetrating or disturbing the surface of the agar (see Note 14). 5. Allow culture to dry (~10 min) until liquid is absorbed into agar (see Note 8). 6. Carefully move the inoculated plates to a 30 °C incubator (see Notes 10 and 15). Keep plates level, do not invert prior to incubation (see Note 11), and do not stack plates (see Note 12). 7. Monitor the plates (see Note 16); after several hours, migration will be observable around the inoculation site. Over time, the bacteria will move outward across the agar surface (see Note 17) and eventually encounter the border and antibiotic compartment. Further expansion will depend on the strain, experimental conditions, and the agent in use (see Note 8). The time required for incubation will be dependent on the strain of interest as well as the established plate/environmental conditions (see Note 15). 8. Record the colony diameter at the widest point from the border with a ruler and the time taken to establish the colony (see Notes 18 and 21). 9. Image the plates (see Note 6, and Fig. 1c for an example).

4

Notes 1. Here, wild-type Salmonella 14028 is used. Other Salmonella isolates will show similar motility patterns. Protocol details such as culture volume, culture density, and length of incubation can be adjusted to maximize motility of specific isolates (see below). 2. For Salmonella, LB (or other rich media) is commonly used for cultivation and motility assays. Swimming can be evaluated on defined minimal media as well. Modification of growth media will require re-optimization of inoculum size, incubation length, etc. 3. For preparation of soft agar for chemotaxis assays, any commercially available agar is suitable.

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4. When cooling, it is best to utilize a hot plate/stir bar combination to ensure the agar is kept in motion and does not cool below 55 °C, which may otherwise result in partial solidification of the media. 5. Once set, soft agar plates can be used immediately. Storage over a period of time can lead to loss of moisture and inconsistent results. 6. For quality image capture of motility plates, the simple “bucket of light” device is very effective [45]. More hi-tech options are also available, some of which can be used to capture time-lapse images of developing motility assays [34, 38, 46]. 7. Temperate swarmers can be fastidious when it comes to growth conditions. For example, E. coli and Salmonella will not swarm on media containing only minimal salts and need addition of casamino acids. E. coli will only swarm when the media is solidified with Eiken agar (Eiken Chemical, Japan), the texture of which likely supports a more hydrated surface [47, 48]. This agar also better supports Salmonella swarming in comparison to more common commercial brands. Even on rich media, E. coli and Salmonella require supplementation with glucose for swarming to occur [27, 47, 49]. 8. For swarm assays, it is crucial that plates are prepared correctly. If they are too dry, swarming may not initiate [22, 27, 33]. Typically, pouring plates and leaving at room temperature for no more than ~16 h is recommended. Avoid proximity to lit Bunsen burners, direct sunlight, or drying near AC vents. If the plates are too wet, bacteria are more likely to spread passively, as they grow, divide, and push out across the surface [22, 27, 46, 50] (see Note 9). Do not use plates immediately after pouring when they may be too wet. Optimization of swarm assays in a lab may take an amount of trial and error to find prep conditions that are suitable and yield consistent results. Long-term storage of swarm plates is not recommended due to the importance of hydration for swarming. 9. Confirmation of swarming can be achieved using microscopy. Observation of swarm fronts with a 10× objective or 40× longworking distance objective should reveal the characteristic whirls and flows associated with this collective motion (see videos in refs. [21, 27]). This visual confirmation is strongly recommended when first beginning setup and optimization of any swarming experiments to ensure that you are not observing passive spreading. 10. While Salmonella can grow/swim at 37 °C, we find more consistent results when assays are carried out at 30 °C. The lower temperature also allows swim assays to be compared with swarm assays (see Note 15).

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11. Keep plates flat during incubation to avoid disturbing the soft agar. 12. Stacking plates (i.e., putting one on top of the other) can cause uneven temperature distribution within the incubator and affect consistency of the readout. 13. Swim/chemotaxis plates typically show symmetrical, concentric rings, emanating out from the inoculation point. The diameter of the ring is typically measured for these experiments, although the distance of the outer ring from the inoculation point can also be used. Rate of expansion within a given time can serve as a comparative measure of motility between strains and conditions. 14. Inoculate gently, keeping volume of the inoculum small. Aggressive pipetting may aerosolize the culture and disperse across the surface, resulting in emergence of satellite swarm colonies inoculation points across the surface. Avoid trapped air bubbles in the inoculum, which can perturb the shape of the developing swarm. 15. Salmonella will swarm at both 30 °C and 37 °C degrees, but the higher temperature can lead to premature plate drying. If a swarm assay is expected to last for a long period of time, a humidity chamber incubator is recommended (see Note 16). 16. An important concern when monitoring swarm plates is that repeated removal of the lid will result in the loss of moisture, which will hinder (or eventually halt) swarming progress. Refrain from removing the lid until ready to end the experiment. 17. One of the hallmarks of swarming is a notable lag time, which can vary between species, strains, and environmental/media composition [21, 22, 27, 51]. This is another aspect of swarming that can be monitored through these assays. 18. Unlike swim plates, swarm colonies often expand in a less symmetrical fashion. We typically measure diameter at the widest point. Alternatively, you can record the expansion outward from the inoculation point to the furthest point traveled. 19. Swarming bacteria show distinct adaptations to the environment, not evident in swimmers. The border-crossing assay is a simple yet effective way to naturally prime cells into a swarming state. This approach has shed new light on the enhanced resistance of swarming bacteria to an array of antibiotics [31, 32, 35]. We warn against substituting this natural development of a swarm with swarming mimics such as artificially increasing cell length or cell density [52, 53]. 20. If encountering difficulty using this “drag method,” allow the agar to dry, and then slowly pipette ~100 μL of molten agar

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along the interface between the two compartments to create a border. 21. Encroachment of a swarm onto agar supplemented with a challenge condition is likely to slow motility and in some cases exacerbate the lack of symmetry in a swarm colony (see Note 18). Thus, it is also important to observe overall colonization of the right-hand compartment when judging tolerance of the condition and motility in response to a stress.

Acknowledgments Motility research in our lab is supported by grants from NIGMS (R35 GM118085) and NIAID (R21 AI158295). References 1. Miyata M, Robinson RC, Uyeda TQP et al (2020) Tree of motility – a proposed history of motility systems in the tree of life. Genes Cells 25:6–21 2. Harshey RM (2003) Bacterial motility on a surface: many ways to a common goal. Annu Rev Microbiol 57:249–273 3. Berg HC (2004) E. coli in motion, 1st edn. Springer-Verlag, New York 4. Jarrell KF, McBride MJ (2008) The surprisingly diverse ways that prokaryotes move. Nat Rev Microbiol 6:466–476 5. Duan Q, Zhou M, Zhu L et al (2013) Flagella and bacterial pathogenicity. J Microbiol 53:1–8 6. Chaban B, Hughes HV, Beeby M (2015) The flagellum in bacterial pathogens: for motility and a whole lot more. Semin Cell Dev Biol 46:91–103 7. Haiko J, Westerlund-Wikstrom B (2013) The role of the bacterial flagellum in adhesion and virulence. Biology (Basel) 2:1242–1267 8. Nakamura S, Minamino T (2019) Flagelladriven motility of bacteria. Biomolecules 9:279 9. Berg HC (2003) The rotary motor of bacterial flagella. Annu Rev Biochem 72:19–54 10. Xing J, Bai F, Berry R et al (2006) Torque– speed relationship of the bacterial flagellar motor. Proc Natl Acad Sci 103:1260–1265 11. Zhou J, Lloyd SA, Blair DF (1998) Electrostatic interactions between rotor and stator in the bacterial flagellar motor. Proc Natl Acad Sci 95:6436–6441 12. Fukuoka H, Wada T, Kojima S et al (2009) Sodium-dependent dynamic assembly of

membrane complexes in sodium-driven flagellar motors. Mol Microbiol 71:825–835 13. Brown MT, Delalez NJ, Armitage JP (2011) Protein dynamics and mechanisms controlling the rotational behaviour of the bacterial flagellar motor. Curr Opin Microbiol 14:734–740 14. Parkinson JS, Hazelbauer GL, Falke JJ (2015) Signaling and sensory adaptation in Escherichia coli chemoreceptors: 2015 update. Trends Microbiol 23:257–266 15. Larsen SH, Reader RW, Kort EN et al (1974) Change in direction of flagellar rotation is the basis of the chemotactic response in Escherichia coli. Nature 249:74–77 16. Macnab RM (1977) Bacterial flagella rotating in bundles: a study in helical geometry. Proc Natl Acad Sci 74:221–225 17. Berg HC, Brown DA (1972) Chemotaxis in Escherichia coli analysed by three-dimensional tracking. Nature 239:500–504 18. Macnab RM, Koshland DE Jr (1972) The gradient-sensing mechanism in bacterial chemotaxis. Proc Natl Acad Sci 69:2509–2512 19. Adler J (1966) Chemotaxis in bacteria. Science 153:708–716 20. Mesibov R, Adler J (1972) Chemotaxis toward amino acids in Escherichia coli. J Bacteriol 112: 315–326 21. Partridge JD, Harshey RM (2013) Swarming: flexible roaming plans. J Bacteriol 195:909– 918 22. Kearns DB (2010) A field guide to bacterial swarming motility. Nat Rev Microbiol 8:634– 644

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23. Darnton NC, Turner L, Rojevsky S et al (2010) Dynamics of bacterial swarming. Biophys J 98: 2082–2090 24. Jeckel H, Jelli E, Hartmann R et al (2019) Learning the space-time phase diagram of bacterial swarm expansion. Proc Natl Acad Sci 116:1489–1494 25. Zhang HP, Be’er A, Florin EL et al (2010) Collective motion and density fluctuations in bacterial colonies. Proc Natl Acad Sci 107: 13626–13630 26. Kearns DB, Losick R (2003) Swarming motility in undomesticated Bacillus subtilis. Mol Microbiol 49:581–590 27. Partridge JD (2022) Surveying a swarm: experimental techniques to establish and examine bacterial collective motion. Appl Environ Microbiol 88:e0185321 28. Tian M, Zhang C, Zhang R et al (2021) Collective motion enhances chemotaxis in a two-dimensional bacterial swarm. Biophys J 120:1615–1624 29. Partridge JD, Nhu NTQ, Dufour YS et al (2019) Escherichia coli remodels the chemotaxis pathway for swarming. mBio 10: e00316-19 30. Partridge JD, Nhu NTQ, Dufour YS et al (2020) Tumble suppression is a conserved feature of swarming motility. mBio 11:e01189-20 31. Butler MT, Wang Q, Harshey RM (2010) Cell density and mobility protect swarming bacteria against antibiotics. Proc Natl Acad Sci 107: 3776–3781 32. Bhattacharyya S, Walker DM, Harshey RM (2020) Dead cells release a “necrosignal” that activates antibiotic survival pathways in bacterial swarms. Nat Commun 11:4157 33. Harshey RM, Partridge JD (2015) Shelter in a swarm. J Mol Biol 427:3683–3694 34. Morales-Soto N, Anyan ME, Mattingly AE et al (2015) Preparation, imaging, and quantification of bacterial surface motility assays. J Vis Exp 98:52338 35. Partridge JD, Harshey RM (2020) Investigating flagella-driven motility in Escherichia coli by applying three established techniques in a series. J Vis Exp 159:61364 36. Ho¨lscher T, Dragosˇ A, Gallegos-Monterrosa R et al (2016) Monitoring spatial segregation in surface colonizing microbial populations. J Vis Exp 116:54752 37. Pearson MM (2019) Methods for studying swarming and swimming motility. Methods Mol Biol 2021:15–25 38. Bru JL, Siryaporn A, Høyland-Kroghsbo NM (2020) Time-lapse imaging of bacterial swarms

and the collective stress response. J Vis Exp 159:60915 39. Heering J, Alvarado A, Ringgaard S (2017) Induction of cellular differentiation and single cell imaging of Vibrio parahaemolyticus swimmer and swarmer cells. J Vis Exp 123:55842 40. Sharp PM (1991) Determinants of DNA sequence divergence between Escherichia coli and Salmonella typhimurium: codon usage, map position, and concerted evolution. J Mol Evol 33:23–33 41. Partridge JD, Nieto V, Harshey RM (2015) A new player at the flagellar motor: FliL controls both motor output and bias. mBio 6:02367 42. Harshey RM (2010) Swarming adventures. In: Maloy SM, Casadesus J, Hughes K (eds) The lure of bacterial genetics. American Society for Microbiology, Washington, DC, pp 163–171 43. Adler J (1969) Chemoreceptors in bacteria. Science 166:1588–1597 44. Hazelbauer GL (2012) Bacterial chemotaxis: the early years of molecular studies. Annu Rev Microbiol 66:285–303 45. Parkinson JS (2007) A “bucket of light” for viewing bacterial colonies in soft agar. Methods Enzymol 423:432–435 ´ T (2017) Sliding on the 46. Ho¨lscher T, Kova´cs A surface: bacterial spreading without an active motor. Environ Microbiol 19:2537–2545 47. Harshey RM, Matsuyama T (1994) Dimorphic transition in Escherichia coli and Salmonella typhimurium: surface-induced differentiation into hyperflagellate swarmer cells. Proc Natl Acad Sci 91:8631–8635 48. Toguchi A, Siano M, Burkart M et al (2000) Genetics of swarming motility in Salmonella enterica serovar typhimurium: critical role for lipopolysaccharide. J Bacteriol 182:6308–6321 49. Mattingly AE, Weaver AA, Dimkovikj A et al (2018) Assessing travel conditions: environmental and host influences on bacterial surface motility. J Bacteriol 200:e00014-18 50. Brown II, H€ase CC (2001) Flagellumindependent surface migration of Vibrio cholerae and Escherichia coli. J Bacteriol 183: 3784–3790 51. Copeland MF, Weibel DB (2009) Bacterial swarming: a model system for studying dynamic self-assembly. Soft Matter 5:1174– 1187 52. Swiecicki JM, Sliusarenko O, Weibel DB (2013) From swimming to swarming: Escherichia coli cell motility in two-dimensions. Integr Biol (Camb) 5:1490–1494 53. Colin R, Drescher K, Sourjik V (2019) Chemotactic behaviour of Escherichia coli at high cell density. Nat Commun 10:5329

Chapter 14 Analysis of Adhesion and Surface Motility of a Spirochete Bacterium Shuichi Nakamura, Jun Xu, and Nobuo Koizumi Abstract Spirochetes are Gram-negative bacteria with helical or flat wave morphology and move using flagella residing beneath the outer membrane. Most commonly, flagellated bacteria swim in liquid. Meanwhile, some species of spirochete not only swim but keep moving after adhering to solid surfaces, and such amphibious motility is believed to be significant for pathogenicity. This chapter focuses on the zoonotic spirochete Leptospira and describes the method for measuring the spirochete adhesion and surface motility. Key words Spirochete, Leptospira, Motility, Swimming, Crawling, Optical tweezers, Cultured cell

1

Introduction Spirochete

Bacteria grouped into the phylum Spirochaeta exhibit a spiral or flat wave cell body and possess flagella within the periplasmic space [1]. Some species of spirochetes, e.g., Treponema pallidum (syphilis), Borrelia burgdorferi (Lyme disease), Brachyspira hyodysenteriae (swine dysentery), and Leptospira interrogans (leptospirosis), are clinically significant. Experiments using animal models have shown the attenuation of virulence in motility-deficient mutants [2–4], suggesting that motility is a common virulence factor of spirochetal pathogens [5]. This chapter focuses on the zoonotic spirochete Leptospira and describes biophysical approaches to characterize their adhesion and motility.

1.2 Two-Type Motility of the Spirochete Leptospira: Swimming and Crawling

The genus Leptospira is distinguished from other spirochete genera by a short-pitch cell body and curved cell poles [1]. Leptospira comprises the pathogenic, intermediate, and nonpathogenic species, and the pathogens (pathogenic and intermediate species) cause worldwide zoonosis leptospirosis. The pathogenic leptospires are maintained in the renal tract of the reservoirs and shed upon

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urination to environments. Contact with the contaminated soil and water via injured skin is the most potential entry of leptospires to hosts [6, 7]. Leptospira spp. have two periplasmic flagella (PFs), and the PFs transform both cellular ends into either spiral-shape or hook shape [1, 8]. The cell-end morphology depends on the direction of flagellar rotation; when viewed from the distal end of the flagellar motor, counterclockwise (CCW) and clockwise (CW) rotations make the cell ends into hook-shape and spiral-shape, respectively. Two PFs rotate not only cell ends but also the entire coiled cell body and propel the cell. The cell-body rotation produces thrust for swimming similarly to externally flagellated bacteria such as Escherichia coli in liquids [9, 10]. When attaching to solid surfaces, leptospires are propelled by the interaction between the rotating cell body and the surface, which is called “crawling” motility [11, 12]. The crawling motility of Leptospira was first observed in 1975 [11], and we have recently reported that the surface motility is conferred by the interaction of mobile adhesion molecules residing over the outer membrane with solid surfaces [12]. We have further investigated the crawling motility of pathogenic strains on the cultured kidney cells of various mammalian species and have shown the possibility that persistent crawling results in severe symptoms [13]. This chapter elaborates on the principles for quantifying the adhesivity and the directivity of crawling. We introduce an experiment using a glass-made chamber and cultured animal cells. Concerning the experiment using cultured animal cells, we describe only the case using dog kidney cells. 1.3 Principle 1: Steady-State Adhesion

Bacterial adhesion is the consequence of transition from the floating (swimming) state (Fig. 1). Let [Boff] denote the floating fraction and [Bon] the adhesion fraction, the transition reaction is [Boff] $ [Bon]. These fractions are the ratio of the number of floating or adhering bacterial cells, Boff or Bon, to the total cell number, Ball: [Boff] = Boff/Ball and [Bon] = Bon/Ball. The temporal change of [Bon] is appropriate to track the increment of adhering bacteria quantitatively. When reaching the equilibrium, the reaction

Fig. 1 Adhesion analysis. (a) Schematic time course of bacterial adhesion. (b) Increase in the fraction of adhering bacteria with time

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is steady, where bacterial adhesion and detachment from surfaces balance. In the steady-state, the equilibrium constant, Keq = [Boff]/[Bon], is another value to evaluate the bacterial adhesivity [13, 14]. The equilibrium constant links with the free energy difference between the two states, ΔG, as Keq = exp (-ΔG/kBT). 1.4 Principle 2: Diffusion by Crawling

After adhering to the surface, some bacteria stick without migration, and others move with frequent reorientation or translate smoothly. The mean-square displacement (MSD) is a physical parameter to evaluate the diffusivity of micro-objects, like proteins [15] and microorganisms [13, 14, 16]. Figure 2 explains what you can know by MSD analysis. Assuming two particles exist on x axis, one moves to both positive and negative directions with the same

Fig. 2 Crawling analysis using mean square displacement (MSD) of bacterial movement. (a) Simulated one-dimensional displacements of particles that move to positive and negative directions at the same probability (black), biased to positive (red), and move only to positive (green). (b) MSD vs. time plots obtained by analyzing a. (c) Log-log plots of b. (d) Procedure to make MSD vs. time plot. Swimming trajectory (not actual data) shows bacterial positions determined at an interval of 1/30 s. This example calculates the mean values of square displacements (d2) per 1/30, 2/30, and 3/30 s

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possibility (P+ and P-, P+ + P- = 1), and the other’s motion is biased to the positive side. Automatically generated random numbers (rnd) determine their displacements. For example, if rnd < P+, the particle moves in the positive direction; if rnd > P+, it moves in the negative direction. Figure 2a shows simulated particle displacements. The MSD per unit time Δt is h(xi + Δt - xi)2i, where xi is the particle position at time i. The time vs. MSD plot of the diffusive motion shows linear relation to time, whereas another particle that was assumed to move with a bias in the positive direction shows a quadratic curve (Fig. 2b). The linear and quadratic MSD plots are converted to linear lines with slopes of ~1 and ~2 by doublelogarithmic representation, respectively (Fig. 2c). Thus, the MSD slope represents the diffusivity and directivity of crawling. To obtain MSD- t plot, calculate the values of MSD at the interval of unit time using the time course of x–y positions of a crawling bacterium. Assuming that the bacterial position is determined at an interval of 1/30 s, MSD per 1/30 s is given by h(xi + 1/ 2 2 30 - xi) + (yi + 1/30 - yi) i, which is the average of the squared displacements during 1/30 s (denoted by d1, i2 in Fig. 2d). In the same manner, MSD per 2/30 s is given by h(xi + 2/30 - xi)2 + (yi + 2/ 2 2 30 - yi) i, and MSD per 3/30 s is given by h(xi + 3/30 - xi) + (yi + 3/ 2 30 - yi) i. Plot the MSD values against time (Fig. 2d, right bottom), and determine the slope of a regression line fitted to the MSD vs. time plot.

2

Materials Prepare solutions using special grade chemicals and ultrapure water.

2.1

Bacterial Strain

2.2 Media for Leptospira

1. Leptospira spp. (see Note 1). 1. Enriched Ellinghausen-McCullough-Johnson-Harris (EMJH) liquid medium: Mix 902 mL of EMJH basement with 98 mL of EMJH supplement, pH 7.4. Store at 4 °C. 2. EMJH basement: Dissolve 2.3 g of Difco Leptospira medium base EMJH (Becton Dickinson) in 900 mL of distilled water (see Note 2). Add 1 mL of glycerol (stock 10 g glycerol/ 100 mL distilled water, sterilized at 121 °C for 20 min and stored at 4 °C) and 1 mL of sodium pyruvate (stock 10 g sodium pyruvate/100 mL distilled water, sterilized with 0.22-μm pore-size filter and stored at 4 °C). Sterilize EMJH basement at 121 °C for 20 min, and store at 4 °C. 3. EMJH supplement: Dissolve 10 g of bovine serum albumin in 50 mL of distilled water. Mix 1 mL of ZnSO4 (stock 0.4 g ZnSO4·7H2O/100 mL), 1 mL of MgCl2 (stock 1.5 g MgCl2·6H2O/100 mL), 1 mL of CaCl2 (stock 1.5 g CaCl2·2H2O/100 mL), 12.5 mL of Tween 80 (stock 10 g

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Tween 80/100 mL), 1 mL of cyanocobalamin (stock 0.02 g cyanocobalamin/100 mL), 10 mL of FeSO4 (0.05 g FeSO4·7H2O/10 mL, prepare just before making EMJH supplement), and adjust to 98 mL with sterilized water. Store stock solutions at 4 °C after sterilizing MgCl2, CaCl2, and Tween 80 stocks at 121 °C for 20 min, and ZnSO4 and cyanocobalamin stocks with 0.22-μm pore-size filter. 4. 20 mM sodium phosphate buffer (pH 7.4, sterilize at 121 °C for 20 min) (see Note 3). 2.3 Bacterial Growth and Harvest

1. Incubator. 2. 15 mL centrifugal tube (sterilized). 3. 1.5 mL tube (sterilized). 4. Microcentrifuge.

2.4 Microscope Setup

5. Upright microscope (see Note 4). 6. Dark-field condenser (see Note 5). 7. 40× objective. 8. Video camera (see Note 6).

2.5 Glass-Made Flow Chamber

1. Glass slide (0.8 ~ 1.0 mm in thickness) (see Note 7). 2. Coverslip (0.13 ~ 0.17 mm in thickness). 3. Double-sided tape.

2.6 Slide Chamber Cultivating Kidney Cells

1. Chamber slide (chamber size 10 mm × 20 mm). 2. MDCK-NBL2 (dog kidney epithelial cell). 3. Eagle’s minimum essential medium (MEM). 4. Fetal bovine serum. 5. 5% antibiotic/antimycotic mixed solution. 6. 0.1% (w/v) trypsin—0.02% (w/v) EDTA solution. 7. CO2 incubator.

2.7 Software for Data Analysis

3 3.1

The analyses described in this chapter are performed by ImageJ and Microsoft Excel.

Methods Adhesion Assay

1. Grow Leptospira cells at 30 °C for 4 days in EMJH liquid medium (15 mL centrifugal tube) until the late-exponential phase. 2. Centrifuge 500 μL of Leptospira cells (1.5 mL tube) at 1000 g for 10 min at room temperature.

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3. Remove the supernatant, and add 500 μL of 20 mM sodium phosphate buffer. 4. Resuspend the precipitated bacteria with gentle tapping. 5. Infuse the bacterial suspension into a glass-made flow chamber (see Note 8). 6. Set the flow chamber on a microscope stage. 7. Record bacteria using a CCD camera at 30 frames per second. 8. Analyze the movie using image-analysis software. 3.2 Crawling Assay Using a Glass-Made Flow Chamber

1. Grow Leptospira cells at 30 °C for 4 days in EMJH liquid medium until the late-exponential phase. 2. Centrifuge 500 μL of the Leptospira culture at 1000 g for 10 min at room temperature. 3. Remove the supernatant, and add 500 μL of 20 mM sodium phosphate buffer. 4. Resuspend the precipitated bacteria with gentle tapping. 5. Infuse the bacterial suspension into a flow chamber. 6. Set the flow chamber on a microscope stage. 7. Observe the cells, and record movies on a computer. 8. Analyze the movie using image-analysis software; Fig. 3b shows example data obtained by measuring bacteria moving on a glass slide, as depicted in “Example Setup 1” of Fig. 3a.

3.3 Crawling Assay on Kidney Cells Cultivated in a Chamber Slide

1. Cultivate MDCK cells in Eagle’s minimum essential medium containing 10% (w/v) fetal bovine serum and 5% (w/v) antibiotic/antimycotic mixed solution at 37 °C in the presence of 5% CO2, using a cell cultivation flask (see Note 9). 2. Harvest kidney cells in 0.1% (w/v) trypsin and 0.02% (w/v) EDTA and plate on a chamber slide. 3. Incubate the chamber slide at 37 °C with 5% CO2 until a monolayer is formed (~48 h). 4. Remove non-adherent cells by washing twice with antibioticfree media. 5. Incubate the cells for 2 h at 37 °C with 5% CO2. 6. Subject Leptospira prepared as described in Subheading 3.2 to the kidney cells cultivated in the chamber. 7. Incubate the chamber slide containing kidney cells and Leptospira at 37 °C for 1 h. 8. Detach the chamber from the glass slide, and set only the glass slide, where kidney cells adhere, on a microscope stage (see Note 10). 9. Observe and record Leptospira over kidney cells as described in Subheading 3.2.

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Fig. 3 Crawling analysis of leptospires on the cultured kidney cells. (a) Schematic example setups for crawling assay; Setup 1 and Setup 2 analyze bacteria on glass and cultured cells, respectively. A previous study [13] investigated the Leptospira crawling on the cultured kidney cells. (b) Example data obtained by Setup 2 [13]. The upper data show the crawling trails (left), time courses of displacements (middle), and MSD-time plots of bacteria crawling over long distances with a low-frequency reversal (right); the lower ones show limited traveling by crawling with a high-frequency reversal

4

Notes 1. In previous studies, we used some serovars of the pathogenic species Leptospira interrogans [12, 13, 17, 18]. As for adhesion and motility assays, culture medium, growth condition, and sample preparation are common among Leptospira spp.

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2. The approximate formula of Difco™ Leptospira medium base EMJH is as follows: 1.0 g Na2HPO4, 0.3 g KH2PO4, 1.0 g NaCl, 0.25 g NH4Cl, and 0.005 g thiamine per 1 L (final pH 7.5 ± 0.2). 3. For investigating dynamics over the cultured animal cells, we recommend observing leptospires in culture media appropriate for the animal cells. For example, when observing L. interrogans over NRK-52E (rat kidney epithelial cell) and MDCK-NBL2 (dog kidney epithelial cell), we washed leptospires twice in PBS by centrifugation at 1000 g for 10 min and suspended in Dulbecco’s modified Eagle’s medium and Eagle’s minimum essential medium, respectively [13]. 4. Dark-field microscopy can be used to observe the adhesion and crawling of leptospires on glass [12]. Labeling bacteria with fluorescent proteins or dyes is necessary to observe their dynamics on animal cells (“Example Setup 2” in Fig. 3a). In our previous study, a green fluorescent protein (GFP) was constitutively expressed in each leptospiral strain, and a darkfield microscope equipped with an epi-fluorescent system was used [13]. 5. We recommend an oil condenser to acquire high contrast images, but a dry condenser does not severely interfere with the data quality. 6. The frame rate of recording is a critical factor affecting the accuracy of measurements. The appropriate frame rate generally depends on the migration speed, and 30–60 frames per second are sufficient for tracing leptospiral crawling. 7. Leptospira spp. adhere to and crawl over glass surfaces [11, 12] and animal cells [13]. Coating glass with ani-lipopolysaccharide (LPS) antisera affects crawling [12], suggesting that leptospiral surface dynamics involve outer-membrane components. 8. A flow chamber for observation on glass surfaces (Example Setup 1 in Fig. 3a) is made by attaching a glass slide with the washed coverslip with a double-sided tape (Fig. 4).

Fig. 4 Flow chamber to observe leptospires

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9. Dislodge the cells during each passage process by treating them with 0.1% trypsin–EDTA solution. 10. A commercial chamber slide can be separated into a chamber (upper part) and a glass slide (lower part). Since the height of the chamber becomes an obstacle to set 40× objective on a microscope stage, detachment of the chamber part from the glass slide is required.

Acknowledgments We thank Hajime Tahara for his technical support. This work was supported by the JSPS KAKENHI: 18K07100 for SN, 19K07571 for NK, and 18J10834 for JX. References 1. Nakamura S (2020) Spirochete flagella and motility. Biomolecules 10:550. https://doi. org/10.3390/biom10040550 2. Lambert A, Picardeau M, Haake DA et al (2012) FlaA proteins in Leptospira interrogans are essential for motility and virulence but are not required for formation of the flagellum sheath. Infect Immun 80:2019–2025. https://doi.org/10.1128/IAI.00131-12 3. Wunder EA, Figueira CP, Benaroudj N et al (2016) A novel flagellar sheath protein, FcpA, determines filament coiling, translational motility and virulence for the Leptospira spirochete. Mol Microbiol 101:457–470. https:// doi.org/10.1111/mmi.13403 4. Wunder EAJ, Slamti L, Suwondo DN et al (2018) FcpB is a surface filament protein of the endoflagellum required for the motility of the spirochete Leptospira. Front Cell Infect Microbiol 8. https://doi.org/10.3389/ fcimb.2018.00130 5. Nakamura S (2022) Motility of the zoonotic spirochete Leptospira: insight into association with pathogenicity. Int J Mol Sci 23:1859. https://doi.org/10.3390/ijms23031859 6. Picardeau M (2017) Virulence of the zoonotic agent of leptospirosis: still terra incognita? Nat Rev Microbiol 15:297–307. https://doi.org/ 10.1038/nrmicro.2017.5 ˜ a Moctezuma A (2010) Lep7. Adler B, de la Pen tospira and leptospirosis. Vet Microbiol 140: 287–296. https://doi.org/10.1016/j.vetmic. 2009.03.012 8. Bromley DB, Charon NW (1979) Axial filament involvement in the motility of Leptospira interrogans. J Bacteriol 137:1406–1412

9. Goldstein SF, Charon NW (1990) Multipleexposure photographic analysis of a motile spirochete. Proc Natl Acad Sci U S A 87:4895– 4899 10. Nakamura S, Leshansky A, Magariyama Y et al (2014) Direct measurement of helical cell motion of the spirochete Leptospira. Biophys J 106:47–54. https://doi.org/10.1016/j.bpj. 2013.11.1118 11. Cox PJ, Twigg GI (1974) Leptospiral motility. Nature 250:260–261 12. Tahara H, Takabe K, Sasaki Y et al (2018) The mechanism of two-phase motility in the spirochete Leptospira: swimming and crawling. Sci Adv 4:eaar7975. https://doi.org/10.1126/ sciadv.aar7975 13. Xu J, Koizumi N, Nakamura S (2020) Crawling motility on the host tissue surfaces is associated with the pathogenicity of the zoonotic spirochete Leptospira. Front Microbiol 11: 1886. https://doi.org/10.3389/fmicb.2020. 01886 14. Harman MW, Dunham-Ems SM, Caimano MJ et al (2012) The heterogeneous motility of the Lyme disease spirochete in gelatin mimics dissemination through tissue. Proc Natl Acad Sci U S A 109:3059–3064. https://doi.org/10. 1073/pnas.1114362109 15. Kusumi A, Sako Y, Yamamoto M (1993) Confined lateral diffusion of membrane receptors as studied by single particle tracking (nanovid microscopy). Effects of calcium-induced differentiation in cultured epithelial cells. Biophys J 65:2021–2040. https://doi.org/10.1016/ S0006-3495(93)81253-0

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16. Wu X-L, Libchaber A (2000) Particle diffusion in a quasi-two-dimensional bacterial bath. Phys Rev Lett 84:3017–3020. https://doi.org/10. 1103/PhysRevLett.84.3017 17. Takabe K, Nakamura S, Ashihara M et al (2013) Effect of osmolarity and viscosity on the motility of pathogenic and saprophytic

Leptospira. Microbiol Immunol 57:236–239. https://doi.org/10.1111/1348-0421.12018 18. Takabe K, Tahara H, Islam MS et al (2017) Viscosity-dependent variations in the cell shape and swimming manner of Leptospira. Microbiology 163:153–160. https://doi.org/ 10.1099/mic.0.000420

Chapter 15 Force Measurement of Bacterial Swimming Using Optical Tweezers Keigo Abe, Kyosuke Takabe, and Shuichi Nakamura Abstract Velocity is a physical parameter most commonly used to quantify bacterial swimming. In the steady-state motion at a low Reynolds number, the swimming force can be estimated from the swimming velocity and the drag coefficient based on the assumption that the swimming force balances with the drag force exerted on the bacterium. Though the velocity-force relation provides a significant clue to understand the swimming mechanism, the odd configuration of bacteria could develop problems with the accuracy of the force estimation. This chapter describes the force measurement using optical tweezers. The method uses parameters obtained from the shape and movement of a microsphere attached to the bacteria, improving the quantitativeness of force measurement. Key words Swimming force, Optical tweezers, Spirochete, Leptospira, Motility

1

Introduction

1.1 Force of Bacterial Motion

The form of bacterial motility is diverse: Some species possessing flagella in the cell exterior swim by rotating helical propellers, and others move on solid surfaces using pili, leg-like architectures, and adhesins [1]. Whatever the mode of locomotion is, velocity is the most popular parameter that quantitatively characterizes bacterial motility. The velocity (v) can be measured by determining a displacement (Δx) of the bacterial cell with time (Δt): v = Δx/Δt. The force (F) is another crucial physical parameter, but the force measurement is not as easy as the velocity. In steady-state swimming, the swimming force can be estimated from v and the drag coefficient (γ): F = γv. This estimation is based on an assumption that the swimming force balances the drag force exerted on the bacterium and is expected to ensure the quantitativeness for species with simple morphology, such as spherical and rod shapes, where a

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Fig. 1 Leptospira. (a) Dark-field micrographs. (b) Schematic explanation of the structure of Leptospira cell body. Figure 1 of [2], licensed under CC BY 4.0, was reused

Fig. 2 Schematic view of optical tweezers. (a) When an object is away from the laser focus point by the external force, a restoring force pushes it back toward the center of trap. (b) The force applied to the particle linked with a solid wall via a spring with the spring constant of k is proportional to the extension from the equilibrium length (Δx), following the Hooke’s law: F = kΔx. (c) The response of the trapped object against external force is analogous to that of spring, thus the Hooke’s law applies

hydrodynamic theory can determine γ at relatively high precision. However, bacteria with complicated configurations, such as spiral, wavy, and curvy, could develop problems in the accuracy of the force estimation. 1.2 Force Measurement Using Optical Tweezers

We have recently measured the swimming force of a spirochete, Leptospira (Fig. 1), using optical tweezers [2]. The optical tweezer is an experimental technique where a microscale object (e.g., microbeads and cells) is trapped by a focused infrared laser, allowing us to manipulate the trapped object. When a mobile matter is trapped, the force produced by the movement can be estimated by measuring the displacement of the trapped position (Fig. 2). Force measurement using optical tweezers has revealed that the swimming force of Escherichia coli is ~ 0.6 pN [3]. A gliding bacteria, Mycoplasma mobile, on the other hand produces a much stronger stall force of ~ 25 pN [4]. These results suggest that direct contact with a solid surface can produce a larger force than propelling by interaction with a liquid.

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Since Leptospira spp. have a spiral shape in its entire cell body and curvatures at both cellular ends, it is difficult to estimate the swimming force from v and γ. Therefore, we have trapped a microbead attached to the leptospiral cell surface and measured the swimming force of this odd-shaped bacteria by analyzing the bead movement. Our measurement has shown that leptospires generate ~ 17 pN by swimming in liquid [2]. 1.3 Motility and Pathogenicity

The genus Leptospira comprises pathogenic, intermediate, and saprophytic species, and the pathogenic Leptospira cause leptospirosis, a worldwide zoonosis [5, 6]. Leptospires are maintained in the kidney of reservoirs (typically rodents) or animals recovered from symptoms and are shed with urine into environments. Various mammals will be infected via injured skin by contacting contaminated soil or water. Experiments using animal models have shown that motility-deficient mutants of pathogenic Leptospira are considerably attenuated, suggesting that the leptospiral pathogenicity somehow involves motility [7, 8]. As for the percutaneous infection, a certain level of thrust force given by swimming would be required to penetrate the host skin dermis. The powerful swimming of Leptospira that is ~ 30 times greater than E. coli may be significant for infection [2, 9].

1.4 Force Balance in Trapped Bacterium

When a focused laser traps a bead attached to a swimming bacterium, the relevant forces to the system are balanced (Fig. 3) as follows: F = F trap þ F drag ,

ð1Þ

where F is a swimming force, Ftrap is a trapping force, and Fdrag is a drag force exerted on the bead. In other words, Ftrap is a restoring force of optical tweezer, drawing the bacteria moving away from a focusing point of laser back. A displacement of the trapped bead (Δx) can be calibrated to Ftrap using the equation F trap = kΔx,

ð2Þ

Fig. 3 Forces acting on a trapped bead attached to Leptospira. Symbols are explained in the main text

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Fig. 4 Determination of the spring constant of optical tweezers. (a) 2D scatter plot of the position of a 1 μm bead trapped by optical tweezers. (b) Time course of the bead position on x-axis. (c) The distribution of the positions of the bead (histogram) and the estimated potential profile (circles). The dashed line indicates the results of the curve fitting by the harmonic function

where k is a spring constant of the laser trap and is determined from a positional fluctuation of the trapped bead free from bacteria. The value of Fdrag is determined by Stokes’ law: F drag = 6πμr n˜ v,

ð3Þ

where r is the bead’s radius, v is the migration speed (= swimming speed of the bacterium), and μ is the medium viscosity (see Note 1). 1.5 Principle for the Determination of the Spring Constant of Optical Tweezers

When a bead suspended in a medium is trapped, its position fluctuates within a trapping potential and shows a Gaussian distribution (Fig. 4):
 f ðx Þ / exp - x 2 =2σ 2 , ð4Þ where σ is the standard deviation. The distribution obeys Boltzmann’s law: PðxÞ / expð- ΔU =kB T Þ,

ð5Þ

where ΔU is the potential energy, kB is the Boltzmann constant (1.38 × 10-23 J/K), and T is the absolute temperature (296 K). Comparison of Eq. 5 with Eq. 4 gives

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ΔU = ðkB T =2σ2 Þx 2 :

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ð6Þ

Since the thermal fluctuation of the bead trapped by a spring with the spring constant k can be described by the harmonic function (dashed line in Fig. 4c): U ðx Þ = 1=2kx 2 :

ð7Þ

Comparison of Eq. 7 with Eq. 6 gives k = kB T =σ2 :

ð8Þ

Therefore, the value of k is calculated from σ determined by Gaussian fitting to the positional distribution of the trapped bead. Determination of the spring constant is conducted in each flow chamber by trapping a bead free from bacteria.

2

Materials Prepare solutions using special grade chemicals and ultrapure water.

2.1 Materials for Leptospira Strains

1. Leptospira kobayashi strain E30 and Leptospira biflexa strain Patoc I. 2. Enriched Ellinghausen-McCullough-Johnson-Harris (EMJH) liquid medium: Mix 902 mL of EMJH basement with 98 mL of EMJH supplement, pH 7.4. Store at 4 °C. 3. EMJH basement: Dissolve 2.3 g of Difco Leptospira medium base EMJH (Becton Dickinson) in 900 mL of distilled water (see Note 2). Add 1 mL of glycerol (stock 10 g glycerol/ 100 mL water, sterilized at 121 °C for 20 min, and stored at 4 °C) and 1 mL of sodium pyruvate (stock 10 g sodium pyruvate/100 mL water, sterilized with 0.22 μm pore-size filter and stored at 4 °C). Sterilize EMJH basement at 121 °C for 20 min, and store at 4 °C. 4. EMJH supplement: Dissolve 10 g of bovine serum albumin in 50 mL of water. Mix 1 mL of ZnSO4 (stock 0.4 g ZnSO4·7H2O/100 mL), 1 mL of MgCl2 (stock 1.5 g MgCl2·6H2O/100 mL), 1 mL of CaCl2 (stock 1.5 g CaCl2·2H2O/100 mL), 12.5 mL of Tween 80 (stock 10 g Tween 80/100 mL), 1 mL of cyanocobalamin (stock 0.02 g cyanocobalamin/100 mL), and 10 mL of FeSO4 (0.05 g FeSO4·7H2O/10 mL, prepare just before making EMJH supplement), and adjust to 98 mL with sterilized water. Store stock solutions at 4 °C after sterilizing MgCl2, CaCl2, and Tween 80 stocks at 121 °C for 20 min, and ZnSO4 and cyanocobalamin stocks with 0.22 μm pore-size filter.

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Fig. 5 Optical tweezer system. Figure 3a shown in [2], licensed under CC BY 4.0, was reused

5. 20 mM sodium phosphate buffer (pH 7.4, sterilize at 121 °C for 20 min). 6. Incubator. 7. 15 mL centrifuge tube (sterilized). 8. 1.5 mL tube (sterilized). 9. Microcentrifuge. 2.2 Microscope Setup for Optical Tweezers

1. Dark-field microscope (see Note 3) (Fig. 5). 2. 1064 nm semiconductor laser. 3. ×100 oil immersion objective lens (see Note 3). 4. Video camera (see Note 4).

2.3 Materials for Bead Labeling

1. φ1.0 μm carboxyl latex beads (see Note 5).

2.4

1. Glass slide (0.8 ~ 1.0 mm in thickness).

Flow Chamber

2. Coverslip (0.13 ~ 0.17 mm in thickness). 3. Double-sided tape.

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3.1 Preparation of Bacterial Sample

1. Grow Leptospira cells at 30 °C for 4 days in EMJH liquid medium (15 mL centrifugal tube) until the late-exponential phase. 2. Centrifuge 200 μL of bacterial culture (1.5 mL tube) at 1000 g for 10 min at 23 °C. 3. Suspend the cell pellet into a motility medium without pipetting and dilution. 4. Dilute beads with 20 mM sodium phosphate buffer (1:25). 5. Add 10 μL of bead solution to the bacterial suspension, and incubate at 23 °C for 10 min. 6. Make a flow chamber by attaching a glass slide to a coverslip with double-sided tape (Fig. 6). 7. Infuse the bacteria-bead mixture into the flow chamber.

3.2 Measurement of Spring Constant

1. Trap a bead free from bacteria. 2. Record the trapped bead, and analyze the positional distribution. 3. Determine σ by fitting a Gaussian function to the positional distribution of the trapped bead. 4. Calculate k from σ using Eq. 8 (see Note 6).

3.3

Δx-F Calibration

1. Place a flow chamber on a microscope stage. 2. Search bacteria swimming while labeled with a single bead. 3. Migrate the bacteria to the position where a laser is focused by moving the microscope stage, and trap the bead attached to the bacterial surface.

Fig. 6 Flow chamber. A solution (orange) penetrates the space between the coverslip and glass slide by capillary action

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Fig. 7 Example time course of force generation by swimming. The kymograph shows that Leptospira stayed at the laser focus point and swam to the right until the swimming force reached ca. 30 pN. Then, the cell was pulled back to the initial position. The force was calculated from the bead displacement and the swimming speed, as described in Subheading 3.3. Figure 3d of [2], licensed under CC BY 4.0, was reused with modifications

4. Record the bead movement by the bacterial swimming, and analyze the time course of the bead displacement (Δx) (see Note 7). 5. Convert Δx to the swimming force (F) using Eqs. 1, 2, and 3 (Fig. 7).

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Notes 1. The viscosity of water is ~ 0.9 mPa × s at room temperature. A viscometer is available to measure viscosity when an accurate value is required or when viscous agents are supplemented. We used a tuning-fork-type viscometer in our previous study [2]. 2. The approximate formula of Difco™ Leptospira medium base EMJH is as follows: 1.0 g Na2HPO4, 0.3 g KH2PO4, 1.0 g NaCl, 0.25 g NH4Cl, and 0.005 g thiamine per 1 L (final pH 7.5 ± 0.2). 3. Dark-field microscopy is indispensable for observing spirochetes due to their thin body smaller than the optical limit. In principle, the numerical aperture (NA) of the objective for dark-field observation must be lower than that of the condenser lens. However, optical tweezer demands an objective with NA as high as possible for strong trapping. Therefore, you should simultaneously perform dark-field observation and laser trapping using an objective lens with an NA-adjustment ring. 4. The frame rate of recording is a critical factor affecting the accuracy of measurements. The appropriate frame rate generally depends on the thermal fluctuation of beads and the swimming speed of bacteria. We recorded the bead movement at 250 Hz using a COMS (complementary metal-oxide semiconductor) video camera [2]. 5. Optical tweezers can trap φ 0.5 μm beads, but a trapping force is insufficient to measure the Leptospira swimming, i.e., the leptospiral swimming force is larger than the trapping force. Polystyrene beads spontaneously attach to the cell surface of Leptospira [2, 10]. When the bead labeling to specific targets on the cell surface is required, you can conjugate antibodies to the bead surface using the MES buffer, Tris–HCl buffer, 1-(3-dimethylaminopropyl)-3-ethylcarbodiimide (EDAC), and storage buffer, as described in [10]. 6. A spring constant (k) depends on the laser power. Figure 8 shows a relationship between the laser power and k determined by the method described in Subheading 3.2. A value of k affects the resolutions of bead displacement (δx) and force (δF). Namely, δx depends on the thermal fluctuation pffiffiffiffiffiffiffiffiffiffiffiffiffiffi of the trapped bead, 1/2kδx = 1/2kBT, giving δx = kB T =k and F trap = pffiffiffiffiffiffiffiffiffiffiffi kδx = kB Tk. These indicate that the decrease in k increases δx (i.e., lowers the determinant accuracy for bead positions) but decreases δF (improves the force resolution). Since too weak trap releases bacteria, the spring constant should be adjusted to a relatively small value but to an extent that the trapped bacterium does not escape. In [2], we experimented with k = 1636 pN/μm.

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Fig. 8 The spring constant K as a function of the trapping laser power P

7. You may use any software, either commercial or custom-built, as long as it can determine the displacement of the bead position. Data shown in Fig. 6 were analyzed using ImageJ and Microsoft Excel.

Acknowledgments The authors thank Shoichi Toyabe and Jun Xu for their helpful comments and technical supports. This work was supported by the JSPS KAKENHI (18K07100 for SN, 16J01961 for KT). References 1. Miyata M, Robinson RC, Uyeda TQP et al (2020) Tree of motility – a proposed history of motility systems in the tree of life. Genes Cells 25:6–21. https://doi.org/10.1111/gtc. 12737 2. Abe K, Kuribayashi T, Takabe K, Nakamura S (2020) Implications of back-and-forth motion and powerful propulsion for spirochetal invasion. Sci Rep 10:13937. https://doi.org/10. 1038/s41598-020-70897-z 3. Chattopadhyay S, Moldovan R, Yeung C, Wu XL (2006) Swimming efficiency of bacterium Escherichia coli. Proc Natl Acad Sci U S A 103: 13712–13717. https://doi.org/10.1073/ pnas.0602043103 4. Miyata M, Ryu WS, Berg HC (2002) Force and velocity of mycoplasma mobile gliding. J

Bacteriol 184:1827–1831. https://doi.org/ 10.1128/jb.184.7.1827-1831.2002 5. Picardeau M (2017) Virulence of the zoonotic agent of leptospirosis: still terra incognita? Nat Rev Microbiol 15:297–307. https://doi.org/ 10.1038/nrmicro.2017.5 ˜ a Moctezuma A (2010) Lep6. Adler B, de la Pen tospira and leptospirosis. Vet Microbiol 140: 287–296. https://doi.org/10.1016/j.vetmic. 2009.03.012 7. Lambert A, Picardeau M, Haake DA et al (2012) FlaA proteins in Leptospira interrogans are essential for motility and virulence but are not required for formation of the flagellum sheath. Infect Immun 80:2019–2025. https://doi.org/10.1128/IAI.00131-12

Measurement of Bacterial Swimming Force 8. Wunder EA, Figueira CP, Benaroudj N et al (2016) A novel flagellar sheath protein, FcpA, determines filament coiling, translational motility and virulence for the Leptospira spirochete. Mol Microbiol 101:457–470. https:// doi.org/10.1111/mmi.13403 9. Nakamura S (2022) Motility of the zoonotic spirochete Leptospira: insight into association

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with pathogenicity. Int J Mol Sci 23:1859. https://doi.org/10.3390/ijms23031859 10. Tahara H, Takabe K, Sasaki Y et al (2018) The mechanism of two-phase motility in the spirochete Leptospira: swimming and crawling. Sci Adv 4:eaar7975. https://doi.org/10.1126/ sciadv.aar7975

Part III Archaella-Driven Motility of Archaea

Chapter 16 Archaella Isolation Shamphavi Sivabalasarma, Joa˜o N. de Sousa Machado, Sonja-Verena Albers, and Ken F. Jarrell Abstract Swimming archaea are propelled by a filamentous structure called the archaellum. The first step for the structural characterization of this filament is its isolation. Here we provide various methods that allow for the isolation of archaella filaments from well-studied archaeal model organisms. Archaella filaments have been successfully extracted from organisms belonging to different archaeal phyla, e.g., euryarchaeal methanogens such as Methanococcus voltae, and crenarchaeal hyperthermoacidophiles like Sulfolobus acidocaldarius. The filament isolation protocols that we provide in this chapter follow one of two strategies: either the filaments are sheared or extracted from whole cells by detergent extraction, prior to further final purification by centrifugation methods. Key words Archaea, Cell motility, Methanogens, Sulfolobus, Cell surface appendages, Type four filament

1

Introduction Motile archaeal cells are propelled in liquid medium by archaella [1]. This rotating appendage is formed by a long, helical filament, which is stably attached to a membrane-embedded motor. The archaellum motor complex energizes the filament assembly and rotation processes, both powered by ATP hydrolysis [2, 3]. The archaellum is classified as a type four filament (TFF), a group that includes type IV pili (T4P) and type II secretion systems (T2SS), among others [4, 5]. As the only representative of the TFF able to generate torque, the archaellum represents an evolutionarily and mechanistically interesting system. The archaellum filament is formed by archaellins. Archaellins are synthesized as pre-proteins, with a class III signal peptide that must be cleaved prior to archaellin insertion in the growing filament [6–8]. Archaellins are typically modified with N- and, sometimes, O-glycosylation [9, 10]. Most archaellated species encode for multiple archaellin proteins, but the

Tohru Minamino et al. (eds.), Bacterial and Archaeal Motility, Methods in Molecular Biology, vol. 2646, https://doi.org/10.1007/978-1-0716-3060-0_16, © The Author(s), under exclusive license to Springer Science+Business Media, LLC, part of Springer Nature 2023

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function of the different archaellins is not fully understood. In species expressing multiple archaellins, there seems to be no function redundancy, as deletion of one of the archaellins either abolishes or impairs motility [11, 12]. In some cases, different archaellins have been deemed ecoparalogs, i.e., different sets of archaellins are adapted to specific environmental conditions [13, 14]. In other species, it has been suggested that some archaellins are major archaellins, while other archaellins are minor components of the filament enriched in specific regions [15, 16]. Despite biochemical evidence for the existence of different types of archaellins in archaella filaments [10, 17], most of the solved filament structures show a single filament-forming subunit [9, 18, 19]. The only exception so far is Methanocaldococcus villosus, which exhibits a heteropolymeric filament [20]. In this chapter, we provide an overview of methods that have been successfully used to isolate archaella from different organisms. Broadly speaking, these methods consist of variations of two protocols: shearing of cells to remove appendages and detergent extraction from cells. Archaella may also be recovered from spent medium, which can be regarded as a variation of the shearing protocol, without the need to actively force appendage breakage from the cell surface. In this method, archaella filaments have been either recovered directly by centrifugation from the spent medium or concentrated spent medium or precipitated with polyethylene glycol before their recovery [17, 21–23]. It is important to realize that archaellation is non-constitutive in many cases. Archaellation can be influenced by factors like growth medium, temperature, and growth phase [23–27]. Thus, before setting up an archaella isolation experiment, it is necessary to ensure that cells are synthesizing archaella in the first place, either by electron microscopy or motility assays.

2

Materials Ultrapure water (18 MΩ -cm at 25 °C) is used for the preparation of buffers and media.

2.1 Isolation of Archaella from Sulfolobus acidocaldarius by Shearing with a Syringe Needle

1. Sulfolobus acidocaldarius. 2. Brock I (1000× stock solution): Dissolve CaCl2·2H2O (70 g/L) in water and autoclave. 3. Brock II (100× stock solution): Dissolve (NH4)2SO4 (130 g/L), MgSO4·7H2O (25 g/L), KHPO4 (28 g/L) in water. Add 50 mL of trace element solution (see above), adjust to pH 3 with 98% H2SO4, and autoclave it. 4. FeCl3 solution (1000× stock solution): Dissolve 20 g/L FeCl3·6H2O, and filter-sterilize the solution.

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5. NZ-amine: Prepare a 20% (w/v) NZ-amine stock solution and autoclave. 6. Dextrin: Prepare a 20% (w/v) Dextrin stock solution and autoclave. 7. Uracil: Prepare a 5 mg/mL stock solution in warm, ultrapure water. Once dissolved, filter-sterilize and store it at -20 °C. 8. H2SO4. 9. 0.5 g/mL CsCl (ρ = 1.36 g/L). 10. Incubator (75 °C). 11. Spectrophotometer. 12. Floor-standing centrifuge with rotor (e.g., Beckman Coulter centrifuge equipped with a JLA 8.1 rotor, Sorvall RC+ centrifuge equipped with a SS-34 rotor). 13. Floor-standing ultracentrifuge with rotor (e.g., Beckman Coulter ultracentrifuge L-60 equipped with the 60Ti rotor). 14. Desktop ultracentrifuge with rotor (e.g., Beckman Coulter Optima MAX-XP ultracentrifuge with a MLA-55 rotor, a MSL-50 swing out rotor or a MLA-80 rotor). 15. Centrifugation tubes. 16. Ultracentrifugation tubes. 17. Peristaltic pump. 18. Syringe needle with 1.1 mm in diameter and 0.4–0.5 mm in diameter. 2.2 Shearing Archaella from Methanogens with a Waring Blender

1. Strains: Methanococcus, Methanothermococcus, Methanogenium, and Methanoculleus species. 2. Mineral solution 3 for Balch medium III [28]: Dissolve KCl (0.67 g/L), MgCl2·2H2O (5.5 g/L), MgSO4·7H2O (6.9 g/ L), NH4Cl (0.5 g/L), CaCl2·2H2O (0.28 g/L), and K2HPO4 (0.28 g/L) in 1 L of water. 3. FeSO4·7H2O: Dissolve 0.2 g/100 mL water. Add three drops of concentrated HCl. 4. Rezasurin (redox indicator): Dissolve 0.1 g/100 mL water. 5. Sodium carbonate (Na2CO3): Dissolve 8 g/100 mL water. 6. Trace elements (100× stock solution): Dissolve nitrilotriacetic acid (1.5 g/L) in 900 mL water, and adjust to pH 6.5 with 3 N KOH. Add MgSO4·7H2O (3 g/L), MnSO4·2H2O (0.5 g/L), CoSO4 or CoCl2 (0.1 g/L), ZnSO4 (0.1 g/L), CuSO4·5H2O (0.01 g/L), AlK(SO4)2 (0.01 g/L), H3BO3 (0.01 g/L), NiCl2·6H2O (0.025 g/L), Na2SeO3·5H2O, and Na2MoO4·2H2O (0.01 g/L). Finally adjust the pH to 7 with 3 N KOH, and adjust final volume to 1 L water. Trace vitamins

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(100× stock solution): Dissolve biotin (2 mg/L), folic acid (2 mg/L), pyridoxine-HCl (10 mg/L), thiamine-HCl (5 mg/L), riboflavin (5 mg/L), nicotinic acid (5 mg/L), DL-calcium pantothenate (5 mg/L), vitamin B12 (0.1 mg/ L), p-aminobenzoic acid (5 mg/L), and lipoic acid (5 mg/L) in 1 L water. 7. TRIS buffer: 20 mM Tris–HCl, pH 7.5 (add 2% (w/v) NaCl when working with osmotically fragile cells). 8. HEPES buffer: 20 mM HEPES-NaOH, pH 7.5 (add 2% (w/v) NaCl when working with osmotically fragile cells). 9. KBr: Dissolve .0.5 g KBr per each 1 mL of TRIS or HEPES buffer with or without 2% (w/v) NaCl. 10. Shaker. 11. Spectrophotometer. 12. High-speed centrifuge with rotor (e.g., Sorvall RC-2B centrifuge equipped with a GAS-6 rotor). 13. Floor-standing ultracentrifuge with rotor (e.g., Beckman Coulter L8-70M ultracentrifuge with a SW28 or SW41 rotor). 14. Centrifugation tubes. 15. Ultracentrifugation tubes. 16. Waring Blender (model 5011, Baxter Corp, Canlab Division) or equivalent. 2.3 Isolation of Archaella by Detergent Extraction of Whole Cells of Methanogens

1. Media component, centrifuges, and tubes are the same as those described in Subheading 2.2. 2. Polyethylene glycol (PEG) 8000. 3. DNase. 4. RNase. 5. Nonionic detergent OP-10 (Nikko Chemicals Co. Tokyo, Japan). 6. Precipitation buffer: 1 M NaCl, 20% (w/v) PEG 8000. 7. 50 mM Tris–HCl or 50 mM HEPES buffer, pH 7.5 (add 2% (w/v) NaCl when archaella will be prepared from osmotically fragile cells).

2.4 Observation of Archaella by Transmission Electron Microscopy

1. Citrate buffer for archaella filaments from S. acidocaldarius: 25 mM sodium citrate/citric acid, 150 mM NaCl, pH 3.5. 2. TRIS or HEPES buffer for archaella filaments from methanogens: 50 mM Tris–HCl or 50 mM HEPES buffer, pH 7.5 (add 2% (w/v) NaCl when archaella will be prepared from osmotically fragile cells). 3. 300 mesh carbon, formvar-coated copper grids freshly glowdischarged.

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4. 2% uranyl acetate. 5. Filter papers. 6. Transmission electron microscope equipped with either a CCD or CMOS camera.

3

Methods

3.1 Isolation of Archaella by Shearing with a Syringe Needle (See Note 1)

1. Brock medium [29] for S. acidocaldarius growth is prepared from two stock solutions, named Brock I and Brock II, FeCl3, NZ-amine, dextrin, and, for auxotrophic strains, uracil. Preparation of Brock II requires a solution of trace elements. Brock medium is prepared by autoclaving 1 L of water. Add 1 mL of Brock I and 10 mL of Brock II. Add 1 mL of the FeCl3 solution. At this point, a white precipitate may become visible. Supplement with 0.1% (w/v) NZ-amine and 0.2% (w/v) dextrin. When working with uracil auxotrophic strains (all strains derived from MW001 [30]), add uracil at a final concentration of 10 μg/mL. Adjust medium to pH 3.0 with H2SO4. 2. Inoculate S. acidocaldarius (MW156 or MW001) from cryogenic stocks in 5 mL Brock’s medium (see Note 2). Grow the cells at 75 °C with soft agitation for 3–4 days. 3. Inoculate all the pre-culture in 2 L of Brock’s medium. Grow the culture for 48 h until they reach an OD600 = 1. 4. Cool down the cultures to room temperature (RT). 5. Harvest the cells at 5000 × g for 25 min at 4 °C. 6. Suspend the cell pellet in 20 mL of Brock’s medium lacking FeCl3. 7. Use a peristaltic pump with a tubing that is connected to a syringe needle with 10 mm in diameter and 40–50 mm in length (Fig. 1). 8. Pump the cells through the tubing and the syringe needle for 45 min at 25 rpm. 9. Switch the syringe needle to a narrower one with 0.4–0.5 mm in diameter and 10 mm in length. 10. Pump the cells through the syringe using the peristaltic pump at 25 rpm speed for 1 h. 11. After shearing, transfer the cells into suitable centrifugation tubes, and harvest the cells by centrifugation (12,000 × g, 25 min, 4 °C). 12. Carefully transfer the supernatant into ultracentrifugation tubes, avoiding any carryover of residual cell pellet. 13. Ultracentrifuge the sample at 200,000 × g for 1.5 h at 4 °C.

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Fig. 1 Setup of shearing procedure for archaella isolation from S. acidocaldarius. Cells are pumped by the peristaltic pump through tubings connected to a syringe needle

14. Discard the supernatant, and resuspend the pellet in 500 μL of Brock’s medium lacking FeCl3. 15. Dissolve 0.5 g/mL CsCl in Brock’s medium lacking FeCl3, and transfer 4.5 mL to a 5 mL ultracentrifuge tube. Load 500 μL of the dissolved pellet on top of the CsCl solution in the tube. 16. Run density gradient centrifugation (250,000 × g, 4 °C, 16 h) using a MLS50 swinging bucket rotor. 17. Carefully place the tube in a holder. A white band in the first third of the CsCl density gradient should have appeared. 18. Discard the first mL from the top, and then collect the white band (~1 mL). 19. Transfer the white band to an ultracentrifuge tube, and add 4.5 mL of Brock’s medium lacking FeCl3. 20. Ultracentrifuge the isolated filaments at 250,000 × g for 1 h at 4 °C to remove excess CsCl. 21. Resuspend the pellet in 150 μL Brock’s medium lacking FeCl3 or in citrate buffer. 22. Store the isolated filaments at 4 °C. 3.2 Shearing Archaella from Methanogens with a Waring Blender (See Note 3)

1. Add 500 mL of mineral solution 3, 10 mL of trace elements solution (100× stock solution), and 10 mL of trace vitamin solution (100× stock solution) to a 480 mL volume of distilled water. Afterward, add NaCl (18 g/L), FeSO4·7H2O (10 ml stock solution), sodium acetate (1 g/L), yeast extract (2 g/L) (Difco), and trypticase (2 g/L) (BBL) and 1 ml of resazurin solution.

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2. Boil the medium under CO2/N2 (20%/80%) gas mixture, and add sodium carbonate solution (4.8 ml) and L-cysteine-hydrochloride·H2O (0.5 g/L) as well as Na2S·9H2O (0.5 g/L). Solution should turn from blue to pink to colorless as it reduces. 3. Using a glass syringe (30–50 mL size) dispense medium into desired size of serum bottles (120 mL to 1 L sizes) filling to no more than 20% of the bottle volume and crimp with butyl rubber stoppers and aluminum caps. 4. Autoclave serum bottles in a protective container, and after cooling, exchange the headspace gas with CO2/H2 (20%/ 80%) gas mixture to 2 atm (see Note 4). 5. Grow cells in a total of 5 L of Balch medium III at optimum growth temperature by shaking at 150–200 rpm while pressurizing daily with CO2/H2 [20% (v/v) /80%(v/v)]. 6. Harvest cells by centrifugation (8000 × g, 15 min, 15 °C). 7. Resuspend the pellet in 200 mL of TRIS or HEPES buffer supplemented with 2% (w/v) NaCl as an osmoprotectant if required for cell integrity. Resuspension must be gentle to prevent osmotically fragile cells from lysing. 8. Load the resuspended cells in a Waring blender. Volume should be just enough to cover the blades. For the Waring Blender model 5011, this is about 200 mL. 9. Shear the cells at the highest setting for 1 min. Expect foaming after shearing of osmotically fragile cells. You may have to wait a few minutes for the foaming to dissipate. 10. Centrifuge at 8000 × g for 15 min at 15 °C to pellet the cells. 11. Transfer the supernatant containing crude archaella to new centrifuge tubes, and then centrifuge at 20,000 × g for 1 h at 15 °C to remove large membrane fragments arising from the cells broken by shearing. 12. Transfer the supernatant to SW28 ultracentrifuge tubes. Centrifuge at 112,000 × g for 1.5 h to obtain a crude pellet of the archaella. Carefully remove and discard the supernatant. Resuspend the archaella pellet gently in a minimum volume of TRIS or HEPES buffer supplemented with 2% (w/v) NaCl if required for filament integrity (less than 1 mL) (see Note 5). Examination of the pellet at this stage by SDS-PAGE should reveal the archaellins as the major protein bands, typically in the 25–35 kDa range (see Note 6). 13. Pour 12 mL of the KBr solution into SW41 ultracentrifuge tubes.

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14. Apply the crude archaella preparation gently on top of the KBr solution. Make sure that the tubes are filled to the top (see Note 7). 15. Run the density gradient centrifugation (198,000 × g, 20 h, 15 °C). Do not ultracentrifuge at 4 °C as the KBr will likely precipitate at this lower temperature. 16. The archaella will appear as a white band in the lower third fraction of the tube. 17. Remove the archaella band in a small volume (e.g., 1 mL), and transfer to a new SW28 ultracentrifuge tube. 18. Fill the tube with fresh buffer, and ultracentrifuge at 112,000 × g for 90 min at 4 °C to pellet the purified archaella while removing excess KBr. 19. Resuspend the purified archaella in 0.5–1.0 mL of TRIS or HEPES buffer supplemented with 2% (w/v) NaCl if required for archaella integrity for further analysis (see Note 5). 3.3 Isolation of Archaella by Detergent Extraction of Whole Cells of Methanogens (See Note 8)

1. Grow 5 L of cells at their optimum growth temperature with shaking at 150–200 rpm while pressurizing daily with CO2/H2 (80%/20%). 2. Harvest cells by centrifugation (8000 × g, 15 min, 15 °C). 3. Resuspend the cell pellet in 200 mL of TRIS or HEPES buffer with or without 2% (w/v) NaCl. 4. Add nonionic detergent OP-10 at a final concentration of 1% (v/v) along with DNase and RNase (25 μg/mL each) to reduce viscosity of lysed cells (see Note 9). 5. Incubate the cell suspension in detergent with gentle stirring at RT for 30 min. 6. Centrifuge at 8000 × g for 15 min at 15 °C to remove the cell debris. Carefully transfer the supernatant to a beaker (or a series of small beakers), and put on ice. 7. Add the precipitation buffer to a final concentration of 10% (v/v), and gently shake for 1 h on ice (see Note 10). 8. Centrifuge at 8000 × g for 10 min. 9. Carefully remove and discard the supernatant, and resuspend the pellet in less than 1 mL of TRIS or HEPES buffer. 10. Further purification of the crude sample involves banding of the preparation in a KBr gradient. Start by pouring 12 mL of the KBr solution into SW41 ultracentrifuge tubes. 11. Carefully layer the crude archaella preparation on the top of the KBr solution. Make sure that the tubes are filled to the top (see Note 7).

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12. Centrifuge at 198,000 × g for 20 h at 15 °C. Do not centrifuge at 4 °C as KBr will likely precipitate. 13. The archaella will appear as a white band in the lower third fraction of the tube. Remove the white archaella band in a small volume (e.g., 1 mL), and transfer to a new SW41 tube. 14. Fill the tube with fresh buffer, and ultracentrifuge at 112,000 × g for 90 min at 4 °C to pellet the purified archaella, now free of excess KBr. 15. Resuspend the purified archaella in TRIS or HEPES buffer in less than 1 mL for further analysis. 3.4 Imaging of Negative Stained Archaella by Transmission Electron Microscopy

1. Glow-discharge 300 mesh carbon-/formvar-coated copper grids for 1 min. 2. Apply 5 μL of isolated archaella filaments, and incubate the grid at RT for 1 min. 3. Blot the excess liquid using a filter paper. 4. Apply 20 μL of 2% uranyl acetate, and blot it away. Repeat this step 3 more times. 5. After the final staining step, dry the grid with a filter paper, and store it at RT until usage. 6. Observe negative stained samples by transmission electron microscopy.

4

Notes 1. The methods used to purify archaella from Sulfolobus acidocaldarius can be also used to purify threads and Aap pili [31]. 2. The strain MW156, which is devoid of Aap pili and archaella, can be used for the isolation of thread filaments from S. acidocaldarius. Wild-type strains or MW001 can be used as well [26, 30]. 3. Shearing is used typically on 5 L of cells. Grow cells in Balch medium III from a 10% inoculum to stationary phase. Balch Medium III is typically used for Methanococcus, Methanothermococcus, Methanogenium, and Methanoculleus species. Growth can take 2–3 days or more depending on the species. Shearing is a method used to isolate bacterial flagella, and it should be an acceptable methodology for isolation of archaella from any archaeon. Shearing has also been used for isolation of archaella from Methanospirillum hungatei [32], Thermoplasma volcanium, and Saccharolobus (Sulfolobus) shibatae [33]. Variations on the shearing protocol have also been used to isolate archaella from Pyrococcus furiosus [34] and Natrialba magadii

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[35]. However, we have found that shearing of Halobacterium salinarum leads to excessive foaming such that this method could not be used. It should be noted that most archaella are composed of multiple archaellins and the distribution of these throughout the archaellum itself has rarely been addressed. Shearing could result in an archaella preparation that is deficient in cell surface-proximal archaellins [15]. 4. Bottles are under pressure and may explode upon autoclaving. In our hands, this risk increases if more than 20% of bottle volume is exceeded with medium. Since these methanogens use the CO2/H2 in the headspace as carbon and energy sources, in our hands better growth is obtained filling the serum bottles to only 10–20% of the serum bottle volume. 5. It has been reported that the integrity of archaella filaments of Methanothermococcus thermolithotrophicus is NaCldependent [36]. 6. Analysis of the purified archaella by electron microscopy will reveal short filaments (longer archaella filaments are often sheared to much shorter lengths than what appear on cells) that are typically thinner at 10–14 nm than bacterial flagella (18–24 nm). SDS-PAGE analysis reveals multiple archaellin bands usually in the 25–35 kDa range except for rare instances where only 1 archaellin is present (e.g., Sulfolobales). Figure 2 shows typical micrographs of S. acidocaldarius as well as Methanococcus voltae with corresponding micrographs of the isolated archaella filaments. 7. The banding of crude archaella step has sometimes been done on CsCl gradients [34, 35, 37]. 8. The detergent extraction method has been used on methanogens that have an S-layer as their sole wall component (e.g., Methanococcus). Typically, this method has been used on 5 L of cells. Grow cells in Balch Medium III from a 10% inoculum to stationary phase. Growth can take 2–3 days or more depending on the species. Cell proximal archaella proteins can be enriched by first shearing the cells in a Waring blender to remove most of the cell distal archaella structure. Subsequent collection and detergent extraction from the sheared cells should result in enrichment for cell proximal archaella stubs [15]. 9. A preliminary screening of a number of detergents determined that OP-10 was the most effective in lysing cells, leading to the cleanest archaella preparations with the most anchoring structure (knobs) [15]. Other nonionic detergents may be similarly effective. Attempts to use the OP-10 extraction method were unsuccessful on the thermophilic Methanothermococcus thermolithotrophicus. This species was grown in the same medium as

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Fig. 2 Negative stained electron micrographs of Sulfolobus acidocaldarius and Methanococcus voltae and isolated archaella filaments. (a) Transmission electron micrograph of S. acidocaldarius with various cell surface structures. The archaella are indicated by a red arrow. Scale bar: 500 nm (b) Micrograph of isolated archaella filaments from S. acidocaldarius. Scale bar: 200 nm (c) Electron micrograph of a M. voltae cell with a tuft of 70 archaella filaments. Cell fixed in 4% glutaraldehyde before being negatively stained with 2% phosphotungstic acid (pH 7.0). Scale bar: 1 μm. (Reprinted from Bardy et al. [15]). Fig. 9A. (d) Electron micrograph of detergent extracted archaella from M. voltae (approximately 12 nm in diameter) with visible hooks and polar stubs. The scale bar is 50 nm. (Courtesy of S.–I. Aizawa)

the mesophilic methanogens (Balch medium III), but at a higher temperature of 60 °C. Unfortunately, the procedure always led to a massive gray precipitate which thwarted further purification efforts [38]. 10. We limited our experiments to PEG 8000. We have no data on the use of PEG of other molecular weights in the precipitation step, but PEG 6000 was used successfully in the precipitation step of Haloferax volcanii archaella, for example [37].

Acknowledgments NM, SS, and SVA were supported by the Collaborative Research Centre SFB1381 funded by the Deutsche Forschungsgemeinschaft (DFG, German Research Foundation)—Project-ID 403222702— SFB1381. This study was supported in part by the Excellence

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Initiative of the German Research Foundation (GSC-4, Spemann Graduate School) and in part by the Ministry for Science, Research and Arts of the State of Baden-Wuerttemberg. We would like to thank the EM facility at the Faculty of Biology, University of Freiburg, for access to the TEM for generation of data. The TEM (Hitachi HT7800) was funded by the DFG grant (project number 426849454) and is operated by the University of Freiburg, Faculty of Biology, as a partner unit within the Microscopy and Image Analysis Platform (MIAP) and the Life Imaging Center (LIC), Freiburg. References 1. Albers SV, Jarrell KF (2015) The archaellum: how Archaea swim. Front Microbiol 6:23. https://doi.org/10.3389/fmicb.2015.00023 2. Reindl S, Ghosh A, Williams GJ et al (2013) Insights into FlaI functions in archaeal motor assembly and motility from structures, conformations, and genetics. Mol Cell 49:1069– 1082. https://doi.org/10.1016/j.molcel. 2013.01.014 3. Streif S, Staudinger WF, Marwan W, Oesterhelt D (2008) Flagellar rotation in the archaeon Halobacterium salinarum depends on ATP. J Mol Biol 384:1–8. https://doi.org/10.1016/ j.jmb.2008.08.057 4. Berry J-L, Pelicic V (2015) Exceptionally widespread nanomachines composed of type IV pilins: the prokaryotic Swiss Army knives. FEMS Microbiol Rev 39:1–21. https://doi. org/10.1093/femsre/fuu001 5. Denise R, Abby SS, Rocha EPC (2020) The evolution of protein secretion systems by co-option and tinkering of cellular machineries. Trends Microbiol 28:372–386. https://doi.org/10.1016/j.tim.2020.01.005 6. Albers S-V, Szabo´ Z, Driessen AJM (2003) Archaeal homolog of bacterial type IV prepilin signal peptidases with broad substrate specificity. J Bacteriol 185:3918–3925. https://doi. org/10.1128/JB.185.13.3918-3925.2003 7. Kalmokoff ML, Karnauchow TM, Jarrell KF (1990) Conserved N-terminal sequences in the flagellins of archaebacteria. Biochem Biophys Res Commun 167:154–160. https://doi. org/10.1016/0006-291X(90)91744-D 8. Szabo´ Z, Stahl AO, Albers S-V et al (2007) Identification of diverse archaeal proteins with class III signal peptides cleaved by distinct archaeal prepilin peptidases. J Bacteriol 189: 772–778. https://doi.org/10.1128/JB. 01547-06 9. Poweleit N, Ge P, Nguyen HH et al (2016) CryoEM structure of the Methanospirillum hungatei archaellum reveals structural features

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Chapter 17 Direct Observation of Archaellar Motor Rotation by Single-Molecular Imaging Techniques Yoshiaki Kinosita Abstract Single-molecular techniques have characterized dynamics of molecular motors such as flagellum in bacteria and myosin, kinesin, and dynein in eukaryotes. We can apply these techniques to a motility machine of archaea, namely, the archaellum, composed of a thin helical filament and a rotary motor. Although the size of the motor hinders the characterization of its motor function under a conventional optical microscope, fluorescence-labeling techniques allow us to visualize the architecture and function of the archaellar filaments in real time. Furthermore, a tiny polystyrene bead attached to the filament enables the visualization of motor rotation through the bead rotation and quantification of biophysical properties such as speed and torque produced by the rotary motor imbedded in the cell membrane. In this chapter, I describe the details of the above biophysical method based on an optical microscope. Key words Archaea, Halobacterium salinarum, Haloferax volcanii, Rotary motor, Archaellum, Optical microscopy, Single-molecular techniques, TIRFM, Beads assay, Motile ghost

1

Introduction Motility is seen across all domains of life (Eukaryotes, Bacteria, and Archaea). In Eukaryotes, myosin, kinesin, and dynein play a crucial role in intracellular movement such as muscle contraction and chromosome segregation in cell division, which are all driven by the free energy of ATP hydrolysis [1]. Bacteria show various types of motilities, such as gliding, swimming, and twitching, which are driven by surface appendages composed of supramolecular motility machinery [2]. In archaea, cells show only swimming motility driven by the archaellum (archaeal flagellum), which consists of a reversible rotary motor and a helical filament acting as a propeller (Fig. 1a) [3]. In Euryarchaeota, the archaellum is encoded by two genes for the filaments, arlA and arlB, and by eight genes for the motor complex, arlC to –arlJ [4]. Mutant studies and biochemical data suggest the roles of the ArlC/ArlD/ArlE complex for the

Tohru Minamino et al. (eds.), Bacterial and Archaeal Motility, Methods in Molecular Biology, vol. 2646, https://doi.org/10.1007/978-1-0716-3060-0_17, © The Author(s), under exclusive license to Springer Science+Business Media, LLC, part of Springer Nature 2023

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Fig. 1 Quantification of the structural and functional parameters of helical filaments by fluorescence imaging. (a) Schematics of an archaeal cell with archaella. Motile archaea form archaella, each of which consists of a reversible rotary motor and a helical filament acting as a propeller. The ATP-driven molecular motor rotates in both directions, and the rotation is transmitted to the helical filament on the cell surface to consequently propel the cell forward or backward (see the details for Subheading 1). (b) Schematics of the optical setup (Top). To prevent damage to the objective lens due to laser focusing, the laser power is reduced to 20 mW using ND filters (ND) next to the laser. The beam (green line) is expanded by a factor of 10 using the beam expander (BE). The direction of the laser path can be changed by tilting the mirror M1. The lens outside the microscope should be positioned to be conjugated with the back focal plane of the objective lens (Obj.). If the lens is correctly placed, the laser beam would become the sharpest. The fluorescence from samples illuminated by the green laser (pink dashed line) is transmitted to the dichroic mirror (DM), reflected by mirror M2, and finally captured by the EMCCD camera. Schematics of the objective-type TIRFM is shown at the bottom. The laser beam is focused at the back-focal plane of the objective lens, and the collimated beam is totally reflected at the interface between the glass and water if the incident angle is larger than the critical angle (the refractive angle at 90
). Under the TIRFM, an evanescent field is produced with a 1/e penetration depth of less than 200 nm, and only fluorophores localized in the field are visualized. (c) Fluorescent micrographs taken by epi illumination (left) and TIRF illumination (right). (The figures are reused with permission from Kinosita et al. [13] for (a) with modification and Kinosita and Nishizaka [3] for (c))

switching of motor rotation coupled to the chemotaxis machinery [5], the ArlF/ArlG complex for the interaction with the surface layer (S-layer) [6, 7], ArlF for regulating ArlG filament assembly [7], ArlH for the regulation of the switching between the assembly

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of the archaellar filaments and rotation [8], ArlI as an adenosine triphosphatase (ATPase) essential for both assembly and rotation [9, 10], and ArlJ for the membrane-spanning component. The function and structure of each motor protein have been gradually revealed by genetics and biochemistry. The next step is to reveal how the archaellar motor complex drives rotation and propels the cell body forward. We have recently developed singlemolecular techniques to visualize the rotation of the archaellar motor using a fluorescent dye [3, 11] and a tiny polystyrene bead [12] and have finally characterized biophysical parameters of the archaellar motor such as rotational speed and torque. Additionally, the membrane-permeabilized ghost model has enabled us to control the intracellular chemical conditions and reveal the ATP-coupled motor rotation in the archaellum [13]. In this chapter, currently available methods for biophysical measurements in the archaellar motor are described, which will be also helpful for the measurements of bacterial flagellar motors [14, 15].

2

Materials All buffers are prepared with an ultrapure water (Milli-Q). All chemicals are purchased from commercial suppliers. All experiments were performed at room temperature, unless indicated otherwise.

2.1

Archaea Strains

1. Halobacterium salinarum NRC-1 ATCC 700922 (Hbt. salinarum). 2. Haloferax volcanii RYK28: ΔflgA1ΔpilB3::flgA1 A124C (Hfx. volcanii) (see Note 1).

2.2

Chemicals

1. Bovine serum albumin (BSA). 2. Biotin-NHS ester (e.g., B306; Dojindo). 3. Biotin-PEG2-maleimide (e.g., 21901BID; Thermo Fisher Scientific). 4. Streptavidin-conjugated fluorescent dye (e.g., Dylight 488, 21832; Invitrogen). 5. Fluorescent dye (e.g., PA23001; GE health care). 6. Streptavidin. 7. Desalting column (e.g., NAP5, 17-0853-01; Cytiva). 8. Carboxylated-polystyrene bead (e.g., 200 nm [F6774; Molecular Probes], 500 nm [18720; Polysciences], or 970 nm [PMC 1N; Bangs Laboratory, Inc.]). 9. 1-(3-Dimethylaminopropyl)-3-ethylcarbodiimide Hydrochloride (EDC).

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10. N-Hydroxysulfosuccinimide sodium salt (Sulfo-NHS). 11. Sodium cholate hydrate (e.g., C1254; Sigma Aldrich). 12. DNase. 13. Adenosine triphosphate (ATP). 2.3

Stock Solution

1. 30% (w/v) Saltwater (SW): 50 mM Tris–HCl pH 7.2, 4.1 M NaCl, 150 mM MgCl2, 290 mM MgSO4, 9 mM KCl. 2. 10 Casamino acid solution (Ca-solution): 5% (w/v) casamino acid, pH 7.2 adjusted by 4 N NaOH. 3. 1 M CaCl2. 4. Ca-agar for Hbt. salinarum: 25% (w/v) SW, 0.5% (w/v) Ca-solution, 0.002% (w/v) biotin, 0.005% (w/v) thiamine hydrochloride, 0.01% (w/v) L-tryptophan, and 0.01% (w/v) uracil, 1.0% (w/v) agar (see Note 2). 5. Ca-agar for Hfx. volcanii: 18% (w/v) SW, 0.5% (w/v) Ca-solution, 0.002% (w/v) biotin, 0.005% (w/v) thiamine hydrochloride, 4 mM CaCl2, 1.5% (w/v) agar. 6. Ca-medium: 18% (w/v) SW, 0.5% (w/v) Ca-solution, 0.002% (w/v) biotin, 0.005% (w/v) thiamine hydrochloride, 20 mM CaCl2 (see Note 3). 7. Motility buffer: 20 mM HEPES-KOH, pH 7.2, 1.5 M NaCl, 1 M MgCl2. 8. 1 M MES-KOH, pH 6.1. 9. 1 M potassium phosphate, pH 8.0 (KPi). 10. Bead stock buffer: 10 mM Kpi, pH 8.0, 50 mM KCl, 5% (w/v) Glycerol. 11. Ghost buffer A: 50 mM HEPES-NaOH, pH 7.2, 2.4 M KCl, 0.5 M NaCl, 0.2 M MgCl2, 0.1 M CaCl2 (see Note 4). 12. 1% (w/v) sodium cholate hydrate. 13. Ghost Buffer B: Ghost Buffer A+ 0.03% (v/v) sodium cholate hydrate. 14. Ghost Buffer C: Ghost Buffer A+ 1 mg/mL DNase. 15. 200 mM ATP, pH 7.0 adjusted by 4 N NaOH. 16. Ghost Buffer D: Ghost Buffer A+ ATP.

2.4 Fluorescence Microscope

1. An inverted microscope (e.g., Ti-E; Nikon Instruments) equipped with a 100 objective lens (e.g., Plan Apo TIRF, NA 1.49; Nikon Instruments). 2. Filter set: Dichroic mirror (e.g., ZT532dcrb-UF1; Chroma), emission filter (e.g., NF01-532U; Semrock). 3. Electron multiplying charged coupled device camera (e.g., EMCCD camera, Ixon+ DU860; Andor technology).

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4. Optical table (e.g., RS-2000; Newport). A microscope was fixed on the table. Following materials were fixed on the optical table (see Note 5). 5. Green laser (wavelength of 532 nm; e.g., Compass-315M-100; Coherent). 6. ND filters. 7. Mirror. 8. Lens. 2.5 Phase-Contrast Microscope

1. Upright microscope (e.g., Eclipse Ci; Nikon Instruments) equipped with a 40 objective (Ph, NA 0.75; Nikon Instruments) and a halogen lamp. 2. High-speed scientific complementary metal oxide semiconductor camera (sCMOS camera, e.g., LRH1540N; Digimo) (see Note 6).

2.6

Tunnel Slide

1. Double-sided tape. 2. 22  32 mm glass. 3. 18  18 mm glass.

2.7

Software

1. Image J 1.48v (http://rsb.info.nih.gov/ij/). 2. Andor solis software (Andor technology). 3. Igor Pro. software 8.04 (Hulinks).

3

Methods All experiments were performed at room temperature, unless indicated otherwise. All solutions in a tunnel slide were replaced on a microscope stage. A piece of filter paper was put at the other side to replace the solution in the tunnel slide.

3.1 Cultivation of Hbt. salinarum

1. Thaw 50 mL frozen stock, and streak it onto an agar plate. 2. Incubate at 45
C for 3–5 days until colonies are formed. 3. Scratch a colony, and resuspend it in 1 mL of motility buffer. 4. To remove gas vacuoles, centrifuge the suspension at 7000 g for 2 min (see Note 7). 5. Resuspend the pellet in 1 mL of motility buffer.

3.2 Cultivation of Hfx. volcanii

1. Streak Hfx. volcanii frozen stock onto an agar plate, using a sterilized toothpick. 2. Incubate the plate at 45
C for 4–5 days until colonies are formed.

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3. Scratch a colony, and resuspend it into 5 mL Ca-medium. 4. Incubate the suspension at 42
C for 3 h with shaking at 200 rpm. 5. Transfer 10 μL of culture to 50 mL of Ca-medium. 6. Incubate the cell suspension at 42
C with shaking at 200 rpm until an optical density at 600 nm reaches around 0.1 (see Note 8). 3.3 Preparation of Biotinylated Cells

1. Centrifuge the cultured cells at 7000 g for 3 min, and resuspend the pellet in 1 mL of motility buffer. 2. Add 1 mg/mL biotin-NHS ester for Hbt. salinarum cells and 1 mg/mL biotin-PEG2-maleimide for Hfx. volcanii cells into the above solution. 3. Incubate the cell suspension for 1 h. 4. Centrifuge it at 7000 g for 3 min to remove excess biotin in solution. 5. Discard supernatant, and resuspend the pellet in 1 mL of the motility buffer.

3.4 Preparation of Fluorescent DyeLabeled Cells

1. Mix 0.1 mg/mL streptavidin-conjugated fluorescent dye with the biotinylated cells in a tube (see Note 9). 2. Incubate the cell suspension for 3 min at room temperature. 3. Centrifuge it at 7000 g for 3 min to remove excess fluorescent dye in solution. 4. Discard supernatant, and resuspend the pellet in 1 mL of motility buffer.

3.5 StreptavidinBead Preparation

1. Transfer 500 μL of carboxylated-polystyrene beads (e.g., 500 nm in diameter) to a 1.5 mL microcentrifuge tube. 2. Wash the beads by centrifugation twice to remove surfactant and NaN3 included in commercial stocks using 50 mM MES-KOH, pH 6.1 (see Note 10). 3. Discard the supernatant, and resuspend the pellet in 50 mM MES-NaOH, pH 6.1 containing 2 M EDC and Sulfo-NHS. 4. Incubate the suspension for 1 h at room temperature. 5. Wash the beads by centrifugation three times to remove the free EDC and Sulfo-NHS in solution. 6. Discard supernatant, and resuspend the pellet in 200 μL of 10 mM Kpi containing 1 mg/mL streptavidin. 7. Incubate for 3 h at 4
C with continuous mixing with an endto-end mixer. 8. Wash the beads by centrifugation three times. 9. Discard supernatant, and resuspend the pellet in 1 mL bead stock buffer. 10. Storage the beads at 4
C.

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3.6 Visualization of Swimming Motility of FluorescentLabeled Cells

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All analyses were done using Image J. 1. Infuse motility buffer containing 5 mg/mL BSA into the tunnel slide (see Note 11). 2. Wait for 1 min. 3. Infuse fluorescently labeled cells into the tunnel slide. 4. Excite the fluorescent dye on the surface of cells and archaellar filaments by green laser under epi-illumination (Fig. 1b). 5. Capture the sequential images by an EMCCD camera for 10 s with a temporal resolution of 5 ms. 6. Save tif-images in 12-bit format without compression. 7. Open the image in Image J. 8. Rotate the image so that the cells move linearly in the y direction. 9. Create a rectangle with a width of 2–3 pixels where the cells and flagella are fit (Fig. 2a). 10. To create a kymograph, select the following command (Image!Stacks!Make Montage). The horizontal and vertical axis represents time and distance, respectively (see Note 12). 11. Draw a line at the tip of the cell body, and calculate the distance by multiplying the difference in the number of pixels between the end point and the start point by the scale value (Fig. 2b). 12. The swimming speed is calculated by dividing the distance by the time. 13. The rotary speed of the filament can be obtained by counting the number of propagating waves (Fig. 2b).

3.7 Simultaneous Observation of the Architecture and Function of Helical Filaments Under Total Internal Reflection Fluorescence Microscopy (TIRFM)

1. Infuse fluorescently labeled cells into the tunnel slide without the BSA treatment. 2. Wait for 15 min to allow some ratio of cells stick to the glass surface. 3. Remove unbound cells by motility buffer. 4. Excite the fluorescent dyes on cells by TIRF illumination. 5. Capture the sequential fluorescent images by an EMCCD camera for 20 s with a temporal resolution of 4 ms (Fig. 2c). 6. Save tif-images in 12-bit format without compression. 7. Open the tif-image in Image J. 8. Rotate the image so that the filaments are parallel to the x-axis (Fig. 2c).

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Fig. 2 Quantification of the structural and functional parameters of the archaellum rotation under TIRFM. (a) A TIRF micrograph of a swimming cell. (b) A kymograph produced by the line image extracted from (a) as indicated by the elongated green rectangle. The thick green line in (b) is drawn at the tip of the cell so that the slope of the line directly represents the swimming speed of the cell. Pink dots in the kymograph are placed at the joint between the cell body and the root of the archaella. The number of pink dots per unit time corresponds to the rotary speed of the archaellar filaments. (c) A cell stuck on the glass surface visualized by TIRF illumination. The image intensity profile along the green line in (c) is shown in (d). (e) The time course of intensity change measured in the blue box in (c), from which the speed of archaellum rotation can be measured. The pink line in (c) represents the helical pitch angle of the archaellum. (The figures are reused with permission from Kinosita and Nishizaka [3] for (c–e), with modification)

9. Quantify the filament pitch by drawing a straight line for the above image, pressing Ctrl+K to get a profile of the fluorescence intensity in Image J (green line in Fig. 2c). 10. Fit the intensity profile with a multiple Gaussian to the data (Fig. 2d). 11. Draw a line on the image with the angle tool, and select “Analyze ! Measure” to quantify the pitch angle (pink line in Fig. 2c). 12. The filament radius can be estimated by substituting the quantified value into the following equation: Filament radius (r) ¼ 1/2π pitch ( p)  tan [pitch angle (θ)]. 13. To calculate the rotary speed of the filament, place a 2–3 pixel square on the filament and press Image!Stacks!Plot Z-profile (blue square in Fig. 2c).

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14. The rotary speed is quantified by analyzing the above graph of the change in fluorescence intensity by Fourier transform using Igor pro. Software (Fig. 2e). 3.8 Bead Assay (Fig. 3a)

1. Infuse cells into the tunnel slide. 2. Incubate for 30 min at room temperature. 3. Infuse 20 μL of 500 nm streptavidin-conjugated beads suspended in motility buffer into the tunnel slide. 4. Incubate for 15 min at room temperature to allow the beads to stick to the archaellum. 5. Remove the unbound beads by motility buffer. 6. Capture the phase-contrast images using a sCMOS camera for 30 s at temporal resolution of 10 ms. 7. Save avi-images in 8-bit format without compression. 8. Perform centroid fitting in Igor Pro software to determine the location of ghosts. 9. Convert the x–y position of each frame into the angle (θ) following equation: θ ¼ arctan(y/x). 10. The time course of rotation is produced by dividing the integrated angle by 360. 11. Fit the data by linear fitting to quantify rotary speeds. 12. Estimate the torque generated by the archaellar motor (see Note 13).

3.9 Ghost Preparation

1. Infuse cells into the tunnel slide. 2. Incubate for 30 min. 3. Infuse 20 μL of motility buffer containing 500 nm streptavidin-conjugated beads into the tunnel slide. 4. Incubate for 15 min. 5. To wash unbound beads, infuse 50 μL of the motility buffer into the tunnel slide. 6. Infuse the 25 μL of Ghost buffer B into the tunnel slide. 7. Infuse 25 μL of Ghost buffer C into the tunnel slide when the cell density decreases. 8. To reactivate the archaeal motor, infuse 60 μL Ghost buffer D into the tunnel slide (Fig. 3b, see Note 14). 9. Capture the phase-contrast images using a sCMOS camera for 30 s at temporal resolution of 10 ms. 10. Save images as an avi-file in 8-bit format without compression. 11. Perform centroid fitting in Igor Pro software to determine the location of ghosts (Fig. 3c).

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Fig. 3 Observation of ATP-coupled motor rotation of the archaellum using a ghost cell. (a) Schematics of the experimental setup of bead assay. A cell body is bound to the glass surface to observe behaviors of a single archaellum. The archaellar motor rotation can be observed through the rotation of a streptavidin-decorated bead attached to the biotinylated archaellar filament. (b) The procedure for the ghost-bead assay. Live cells stuck on the glass surface with rotating archaella are permeabilized by a detergent. The change of the cells to ghosts is judged by the reduction of cell density through the phase-contrast images in real time. Subsequently, the detergent is replaced with the buffer with ATP for reactivation of the motor. The phase-contrast images of a live cell (lower Left) and its ghost (lower Right) are shown. (c) Bead movements during ghost preparation. Top: The y-coordinate of the rotating bead during ghost preparation. Bottom: Shaded sections in three different colors in top panel are expanded (left) and their speed distributions are analyzed by Fourier transformation (right), both in the corresponding colors. (d) Left: an x–y plot of the location of a bead attached to an archaellar filament. Upper right: The cumulative angle vs. time for the bead. Lower right: The number of revolutions vs. time for the same data. The red line in the lower right panel represents the linear fitting to the data. Indicating the rotary speed of the archaellar motor. (The figures are reused with permission from Kinosita et al. [13] for (b–d), with modification)

12. Convert the x–y position of each frame into the angle (θ) following equation: θ ¼ arctan(y/x) (Fig. 3d, left panel and upper right panel). 13. The time course of rotation is produced by dividing the integrated angle by 360 (Fig. 3d, lower right panel). 14. Fit the data by linear fitting to quantify rotary speeds (red line in Fig. 3d, lower right panel).

4

Notes 1. Amine-reactive reagents effectively bind to the filaments of Hbt. salinarum [11], whereas this method did not work for the experiments with Hfx volcanii. Therefore, on the environment-exposed surface of the archaellins of Hfx. volcanii, alanine at the 124th amino acid residue was genetically

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replaced with cysteine (ArlA1 A124C) [13]. By mixing this mutant with a maleimide dye (thiol-reactive reagents), the archaellar filaments can be fluorescently labeled. 2. To prevent precipitation, sterilize Ca-medium and SW separately. 3. If the cells are cultured in the concentrations lower than 10 mM CaCl2, the fraction of nonmotile cells increases [13]. 4. Since the S-layer is maintained by CaCl2, more than 10 mM of CaCl2 is required to observe stable rotation [3, 13]. 5. The laser is aligned as shown in ref. [16]. In brief, a lens is used to focus the laser onto the back focal plane of the objective lens. To construct the TIRF illumination, the incident angle of the laser was adjusted by a mirror. 6. For a bead assay, the recording rate should be more than 100 frames/s (10 ms). The maximum speed of the archaeal motor is around 20 Hz, meaning that 50 ms will take for one rotation. The recording rate at 10 ms is sufficient to track the bead movement correctly. 7. Hbt. salinarum forms gas vacuoles in our culture method, which can be easily removed by centrifugation. 8. When the OD at 600 nm exceeds 0.1, the percentage of motile cells gradually decreases [17]. 9. Cy3-streptavidin is prepared by mixing Cy3-NHS-ester and streptavidin. The powder of Cy3 was suspended in 10 mM HEPES-NaOH, pH 7.0 to be 1 mg/mL. After incubation for 1 h at room temperature, the solution was transferred into the NAP5 column to remove the free Cy3 dye. 10. When pelleting beads in a centrifuge, the rotation speed should be adjusted with the bead size: 200 nm beads, 15,000 g for 5 min; 500 nm beads, 12,000 g 5 min for; 1000 nm beads, 10,000 g for 3 min. 11. BSA is needed to prevent cells from adhering to the glass surface. 12. After selecting the command to make montage in Image J, you should type the number of images at columns. The increment and border width should be fixed at 1 and 0, respectively. 13. Use the following equation to calculate the motor torque: T ¼ 2πfξ, where f is rotational speed and ξ ¼ 8πηa3 + 6πηar2 the viscous drag coefficient, with r the radius of rotation of the bead center, a the bead radius, and η the viscosity [18]. 14. The rotation speed depends on the [ATP]. The maximum rotational speed is achieved when 5 mM [ATP] is added, and the Km is approximately 200 μM.

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References 1. Veigel C, Schmidt CF (2011) Moving into the cell: single-molecule studies of molecular motors in complex environments. Nat Rev Mol Cell Biol 12(3):163–176. https://doi.org/10. 1038/nrm3062 2. Miyata M, Robinson RC, Uyeda TQP et al (2020) Tree of motility – a proposed history of motility systems in the tree of life. Genes Cells 25(1):6–21. https://doi.org/10.1111/ gtc.12737 3. Kinosita Y, Nishizaka T (2018) Crosskymography analysis to simultaneously quantify the function and morphology of the archaellum. Biophys Physicobiol 15:121–128. https://doi.org/10.2142/biophysico.15.0_ 121 4. Desmond E, Brochier-Armanet C, Gribaldo S (2007) Phylogenomics of the archaeal flagellum: rare horizontal gene transfer in a unique motility structure. BMC Evol Biol 7:106. https://doi.org/10.1186/1471-2148-7-106 5. Schlesner M, Miller A, Streif S et al (2009) Identification of Archaea-specific chemotaxis proteins which interact with the flagellar apparatus. BMC Microbiol 9:56. https://doi.org/ 10.1186/1471-2180-9-56 6. Banerjee A, Tsai CL, Chaudhury P et al (2015) FlaF is a β-sandwich protein that anchors the archaellum in the archaeal cell envelope by binding the S-layer protein. Structure 23(5): 863–872. https://doi.org/10.1016/j.str. 2015.03.001 7. Tsai CL, Tripp P, Sivabalasarma S et al (2020) The structure of the periplasmic FlaG-FlaF complex and its essential role for archaellar swimming motility. Nat Microbiol 5(1): 216–225. https://doi.org/10.1038/s41564019-0622-3 8. Chaudhury P, Neiner T, D’Imprima E et al (2016) The nucleotide-dependent interaction of FlaH and FlaI is essential for assembly and function of the archaellum motor. Mol Microbiol 99(4):674–685. https://doi.org/10. 1111/mmi.13260 9. Chaudhury P, van der Does C, Albers SV (2018) Characterization of the ATPase FlaI of the motor complex of the Pyrococcus furiosus archaellum and its interactions between the ATP-binding protein FlaH. PeerJ 6:e4984. https://doi.org/10.7717/peerj.4984

10. Reindl S, Ghosh A, Williams GJ et al (2013) Insights into FlaI functions in archaeal motor assembly and motility from structures, conformations, and genetics. Mol Cell 49(6): 1069–1082. https://doi.org/10.1016/j. molcel.2013.01.014 11. Kinosita Y, Uchida N, Nakane D et al (2016) Direct observation of rotation and steps of the archaellum in the swimming halophilic archaeon Halobacterium salinarum. Nat Microbiol 1(11):16148. https://doi.org/10. 1038/nmicrobiol.2016.148 12. Iwata S, Kinosita Y, Uchida N et al (2019) Motor torque measurement of Halobacterium salinarum archaellar suggests a general model for ATP-driven rotary motors. Commun Biol 2:199. https://doi.org/10.1038/s42003019-0422-6 13. Kinosita Y, Mikami N, Li Z et al (2020) Motile ghosts of the halophilic archaeon, Haloferax volcanii. Proc Natl Acad Sci U S A 117(43): 26766–26772. https://doi.org/10.1073/ pnas.2009814117 14. Kinosita Y, Ishida T, Yoshida M et al (2020) Distinct chemotactic behavior in the original Escherichia coli K-12 depending on forwardand-backward swimming, not on run-tumble movements. Sci Rep 10(1):15887. https:// doi.org/10.1038/s41598-020-72429-1 15. Kinosita Y, Kikuchi Y, Mikami N et al (2018) Unforeseen swimming and gliding mode of an insect gut symbiont, Burkholderia sp. RPE64, with wrapping of the flagella around its cell body. ISME J 12(3):838–848. https://doi. org/10.1038/s41396-017-0010-z 16. Nishizaka T, Mizutani K, Masaike T (2007) Single-molecule observation of rotation of F1-ATPase through microbeads. Methods Mol Biol 392:171–181. https://doi.org/10. 1007/978-1-59745-490-2_12 17. Li Z, Kinosita Y, Rodriguez-Franco M et al (2019) Positioning of the motility machinery in halophilic archaea. mBio 10:3. https://doi. org/10.1128/mBio.00377-19 18. Kohori A, Chiwata R, Hossain MD et al (2011) Torque generation in F1-ATPase devoid of the entire amino-terminal helix of the rotor that fills half of the stator orifice. Biophys J 101(1):188–195. https://doi.org/10.1016/j. bpj.2011.05.008

Part IV Type IV-Driven Twitching Motility of Bacteria

Chapter 18 In Situ Structure Determination of Bacterial Surface Nanomachines Using Cryo-Electron Tomography Longsheng Lai, Yee-Wai Cheung, Matthew Martinez, Kathryn Kixmoeller, Leon Palao III, Stefan Steimle, Meng-Chiao Ho, Ben E. Black, Erh-Min Lai, and Yi-Wei Chang Abstract Bacterial surface nanomachines are often refractory to structural determination in their intact form due to their extensive association with the cell envelope preventing them from being properly purified for traditional structural biology methods. Cryo-electron tomography (cryo-ET) is an emerging branch of cryo-electron microscopy that can visualize supramolecular complexes directly inside frozen-hydrated cells in 3D at nanometer resolution, therefore posing a unique capability to study the intact structures of bacterial surface nanomachines in situ and reveal their molecular association with other cellular components. Furthermore, the resolution of cryo-ET is continually improving alongside methodological advancement. Here, using the type IV pilus machine in Myxococcus xanthus as an example, we describe a step-bystep workflow for in situ structure determination including sample preparation and screening, microscope and camera tuning, tilt series acquisition, data processing and tomogram reconstruction, subtomogram averaging, and structural analysis. Key words Bacterial nanomachines, Cryo-electron tomography, Subtomogram averaging, In situ structural biology, Electron microscope alignment, Tilt series acquisition, Cryo-ET grid screening, SerialEM, IMOD, Dynamo

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Introduction Bacterial surface nanomachines are dynamic supramolecular assemblies that participate in diverse activities including cell motility, virulence, biomolecule secretion, surface sensing, and biofilm formation [1–7]. These activities facilitate the engagement of bacteria with their surroundings and promote their survival. Generally, these machineries consist of a cell envelope-spanning core complex termed the basal body and an appendage that protrudes to the extracellular space [8–10]. The type IV pilus machine (T4PM) in the soil-residing predatory bacterium Myxococcus xanthus is such a

Tohru Minamino et al. (eds.), Bacterial and Archaeal Motility, Methods in Molecular Biology, vol. 2646, https://doi.org/10.1007/978-1-0716-3060-0_18, © The Author(s), under exclusive license to Springer Science+Business Media, LLC, part of Springer Nature 2023

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nanomachine that uses a motor (the basal body) to extend a pilus (the appendage) for adhering to surfaces and then retract to move the cell [11]. While biochemical analyses provide general understandings of the function of nanomachines, structures can reveal more mechanistic information such as how their individual components are assembled and interact with one another in 3D to perform functions at molecular and atomic levels. Structural determination is conventionally carried out on purified components by X-ray crystallography, nuclear magnetic resonance spectroscopy, cryoelectron microscopy single particle analysis, or electron crystallography. However, surface nanomachines present a particular challenge to structure determination by these methods due to their complexity, flexibility, and dependency of the native cell envelope as structural supports rendering them difficult to be purified intact. Cryo-electron tomography (cryo-ET), which can elucidate the structure of macromolecular complexes in their native cellular environments (in situ), is a frontier field in structural biology and has seen dramatic advancement in the past decade [12, 13]. In this technique, cells are vitrified on an electron microscopy (EM) grid, and a series of high-magnification 2D images (tilt series) are acquired on a cryo-transmission electron microscope (cryo-TEM) by tilting the sample along the tilt axis every one or few degrees. A 3D reconstruction (tomogram) of the specimen is then produced by back-projecting the 2D images [14]. In addition to cellular samples, cryo-ET can also be applied to purified organelles or macromolecular complexes. In a subsequent process, namely, subtomogram averaging, the individual 3D volumes of the macromolecule of interest in the tomograms are extracted, aligned, and averaged to reduce noise and produce a better-resolved 3D structure [15]. Because of its unique strength, cryo-ET has been used to determine the in situ structures of many bacterial surface nanomachines, such as the T4PMs [11, 16–19], the type II secretion system [20], the type III secretion system [21–24], the flagellar motors [25–32], the Dot/Icm type IVB secretion system [33–36], the Cag type IV secretion system [8, 37], the conjugation system [38], the type V secretion system [39], the type VI secretion system [40–42], and the type IX secretion system [43, 44], leading to a new wave of mechanistic insights into how they work. Taking the T4PM as an example, prior to the cryo-ET studies, structural understandings of the system were limited to individual subunits or subcomplexes, making it infeasible to understand how different components act in concert as a whole nanomachine for function. Using cryo-ET, we were able to determine the ~3 nm resolution in situ structures of the entire T4PM nanomachine as well as its systematic deletion or tagged mutants [11, 18]. These structures allowed us to map the locations of all the ten core components and

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the minor pilins, fit the atomic structures of individual components, and assemble the pseudo-atomic working models of the T4PM, which provide new insights into its mechanism [11, 18]. As a frontier field, the methods and technologies used for cryoET are rapidly evolving, and as a result, there are usually multiple ways to achieve the same goals at each step. In this chapter, we take the M. xanthus T4PM as an example and provide in detail our current protocol for in situ structure determination of bacterial surface nanomachines that includes sample preparation and screening, microscope and camera tuning, tilt series acquisition, data processing, tomogram reconstruction, subtomogram averaging, and structure analysis. This protocol has also been utilized in many other projects in our laboratory such as the rhoptry secretion systems in apicomplexan parasites [45–49].

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Materials

2.1 General Materials

1. M. xanthus wild-type strain DK1622. 2. CTT medium: 1% (w/v) casitone, 10 mM Tris–HCl, pH 7.6, 1 mM KH2PO4, and 8 mM MgSO4. 3. Holey carbon film grids for TEM, R 2/2, 200 mesh, copper support, with extra thickness carbon film, and with or without London Finder reference patterns. 4. Ethane/propane mixture (37% (v/v) ethane and 63% (v/v) propane). 5. Gold colloids, 10 nm. 6. Grade 1 qualitative filter papers, diameter 55 mm. 7. EM grid holder block. 8. AutoGrid C-clip rings. 9. AutoGrid C-clips. 10. Grid storage system, including the storage rack and pucks.

2.2

Instruments

1. AutoGrid assembly workstation and accessories. 2. Autoloader transfer station and accessories. 3. Leica EM GP2 automatic plunge freezer. 4. Leica EM CTD cryo-tools dryer. 5. Glow discharging cleaning system, PELCO easiGlow 91000. 6. Cryo-light microscope. 7. Krios G3i cryo-TEM (Thermo Fisher Scientific). 8. K3 Summit camera (Gatan, Inc.).

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Software

1. TEM User Interface (Thermo Fisher Scientific). 2. Sherpa (Thermo Fisher Scientific). 3. DigitalMicrograph (Gatan, Inc.). 4. SerialEM, version 3.8.6 [50]. 5. IMOD [51]. 6. Dynamo [52]. 7. ChimeraX [53].

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Methods

3.1 Bacterial Culture and Grid Preparation

1. Grow M. xanthus in CTT medium at 32
C with an agitation rate of 150 rpm until the absorbance at 550 nm reaches 0.8. 2. Prepare the gold fiducials by mixing 100 μL of the 10 nm gold colloids with 25 μL of 5% (w/v) bovine serum albumin (BSA) in a 0.5 mL microcentrifuge tube. Vortex for 30 s (s), centrifuge at 15,000  g for 15 min at room temperature, and carefully remove the supernatant. The pellet will be mixed with the bacterial culture in step 10 below (see Note 1). 3. Use a pair of ultrafine tip forceps to grab the grid on the edge and place it on the grid holder block with the dull carbon side facing up. Place the block on the sample table of the PELCO easiGlow 91000 system and sit the glass chamber properly to make sure the O-ring on the base is well positioned. Press the “AUTO RUN” button to start the factory installed program, which will pump to 0.39 mBar, hold for 10 s, glow discharge for 60 s at 15 mA negative polarity and then vent to atmosphere (see Note 2). 4. The sample vitrification on the grids is performed on the EM GP2 automatic plunge freezer. From the main screen, press “Settings.” In the “Environment” panel, set the environment chamber to 32
C, the relative humidity to 85%, “Window heater” to 100%, cryogen temperature to 196
C, and “Cryogen GN2 flow” to 100%. In the “Load Specimen” panel, check “Rotate before specimen application” and uncheck “Rotate to home position after specimen application” if the specimen will be applied from the right-side port. If the specimen will be applied from the left-side port, the option “Rotate to home position after specimen application” should be checked. Set “Delay time before blotting” to 0. 5. In the “Blot” panel, check “single blot,” set the blot time to 6 s or any other time suitable for your sample, and uncheck “Sensor blotting.” Place a Whatman grade 1 filter paper ring onto the magnetic holder and secure it with the magnetic ring (see Note 3).

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6. In the “Plunge/Transfer” panel, check “Automatically plunge after blotting,” set “Post-blotting time” to 0, uncheck “Skip transfer position,” check “Automatically move to transfer position,” and set “Transfer position above freezing position” to 3 mm. 7. Pour liquid nitrogen into the freezing chamber until the “LN2 level” on the screen shows 100%. Place a cryo-grid box into the cryo transfer container and fill it with liquid nitrogen. Place the black secondary cryogen container into its holder. Attach a 1 mL pipet tip to the silicone tubing that is connected to the ethane/propane gas cylinder. Insert the tip into the bottom of the cryogen container and slowly open the valve to allow the ethane/propane mixture to start liquefying. Continue until the container is filled to the brim. 8. Press “Main” and then “LOAD FORCEPS.” Use the manufacturer provided locking forceps to pick up the grid in the desired orientation so that the user can use either the left- or the right-side port for specimen application in step 9 below. Lock the forceps with the locking sleeve and attach it to the gantry below the environment chamber. 9. Mix 15 μL of the bacterial culture with the BSA-coated gold pellet prepared above. The resulting mixture is sufficient for preparing five grids. Press “LOAD SPECIMEN” to lower the environment chamber. From the side port, apply 3 μL of the sample/gold fiducials mixture onto the dull carbon side of the grid. Press “BLOT” to initiate the blotting from the back side of the grid. The forceps will be automatically plunged into the cryogen immediately after blotting. 10. Remove the forceps from the gantry, quickly transfer it to the cryo transfer container, and insert the grid into the assigned slot of the cryo-grid box. 11. Close the cryo-grid box and lock it by tightening the screw. Transfer the box to the puck, insert the puck into the grid storage rack, and store in the cryogenic storage dewar. 3.2 Assembly of AutoGrids

1. The Krios G3i cryo-TEM uses an automatic grid loading system (Autoloader) and requires the grids to be assembled as AutoGrids. AutoGrid refers to the assembly of the C-clip ring, specimen grid and C-clip. Use a pair of ultrafine tip forceps to place the C-clip into the C-clip insertion tool, put the tip of the tool against a flat surface on a paper towel, and press down the plunger. Make sure the C-clip is positioned flat around the inner edge of the tip. 2. Cool the AutoGrid assembly workstation and work in liquid nitrogen vapor. Transfer the AutoGrid box and cryo-grid box into the box-holding slots so that the notch in each box is

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properly locked into position. Place the C-clip ring into its holding position with the flat base facing down and use the long ultrafine tip forceps to transfer the grid from the cryo-grid box onto the C-clip ring with the carbon side facing down. 3. Cool the AutoGrid forceps and the C-clip insertion tool and use either of them to turn and center the ring clipping disk over the grid/C-clip ring assembly. Place the insertion tool on top of the grid, and press down the plunger gently. Turn the disk back, and use the AutoGrid forceps to place the clipped grid assembly in the AutoGrid box with the carbon side facing toward the user. 3.3 Initial Screening of Grids by Cryo-Light Microscope

1. Before loading onto the cryo-TEM, grids can be screened for sample distribution and ice thickness using a cryo-light microscope such as the Leica EM Cryo CLEM described here. Its 50 objective lens allows clear visualization of bacterial cells, while its fluorescence channels allow the localization of molecular assemblies tagged with a fluorescent protein. Furthermore, in the absence of fluorescent tags, the fluorescence channels are useful for assessing the ice thickness due to the autofluorescent properties of the vitrified ice. In Fig. 1, we use grids prepared with Escherichia coli cells to demonstrate these applications. 2. Fill the microscope dewar with liquid nitrogen, and then insert pump into the dewar. Start the pump by pressing “COOL” on the pump control panel. Hold the pump hose near the microscope and wait ~1 min until the nitrogen finishes sputtering and begins to flow smoothly. Next, insert the end of the hose into the stage and lock in place. Wait until the stage reaches 195
C before loading the cartridge onto the stage. The objective should be lowered during cooling so as to properly cool the objective lens. 3. In a humidity-controlled room, cool down the Transfer Shuttle with an AutoGrid cartridge already in place on the transfer arm. Liquid nitrogen should be filled just to the level of the metal grating and no higher. 4. Load AutoGrids into the cartridge with the carbon side facing toward the user. Use the AutoGrid forceps with the thin side away from the user to firmly press the grids into the cartridge slots such that the edges of the C-clip ring are held in place by cartridge springs. Open and slide out forceps, leaving the AutoGrid in place sitting flush to the surface of the cartridge with the carbon side facing up. Slide the protective shield over the loaded AutoGrids and turn the transfer arm to the “closed” position. 5. Carry the Transfer Shuttle to the Leica EM Cryo CLEM and dock it into the microscope. Raise the objective, open the gates between the shuttle and microscope, and slide the cartridge on

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Fig. 1 Cryo-light microscope imaging of E. coli cells on a Quantifoil London Finder grid demonstrating principles which can be applied to M. xanthus or other organisms of interest. (a) A grid square with thick ice unsuitable for cryo-TEM tilt series acquisition imaged by GFP fluorescence (top) and TL-BF (bottom) channels. Note autofluorescence from thick ice throughout the grid square and large cracks through the ice. This grid square was electron opaque by cryo-TEM. (b) A grid square with thinner ice suitable for cryo-TEM tilt series acquisition imaged by GFP fluorescence (top) and TL-BF (bottom) channels. Note the absence of autofluorescence surrounding the target E. coli cell highlighted in red. (c) (Top) The same grid square as in (b), imaged on Krios G3i cryo-TEM at 580 magnification. The target E. coli cell is highlighted in red. (Bottom) 6500 magnification image of the target E. coli cell, demonstrating local ice conditions suitable for cryo-TEM tilt series acquisition

the transfer arm into the microscope. Insert the cartridge onto the stage, turn the transfer arm to “open” position, then retract the arm, and close gates. Lower the objective into the focus position. 6. Open the LAS X software and select desired imaging parameters. Use the transmitted light brightfield channel (TL-BF) as well as the GFP green fluorescence channel to assess ice autofluorescence. If the sample contains any particular fluorophores, add those fluorescence channels as well. 7. Collect a series of images in positions around the grid to assess ice conditions and sample distribution. Montages of the entire grid or areas of the grid can also be collected. 8. The TL-BF channel can be used to assess the overall appearance of the grid square including the presence of contamination. Cracks in the ice as seen by transmitted light indicate thick ice that is generally unsuitable for tilt series acquisition (Fig. 1a).

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9. The fluorescence channel can also be used to assess ice thickness. Thick regions of ice can be identified by looking for autofluorescence. If autofluorescence is present across the full grid square, then the ice is too thick for tilt series acquisition and will not be imageable on TEM (Fig. 1a). Generally, thick ice is seen around the grid bars, but the ice thins as you move toward the center of the grid square. A grid square suitable for TEM imaging will have minimal-to-no autofluorescence in the center of the grid square (Fig. 1b). Use consistent imaging parameters (exposure time, etc.) across grids if wishing to compare their vitrification conditions. 10. Cryo-light microscope images can also be used to identify specific targets for imaging, either based on morphology/positioning or based on fluorescent tagging of specific features, if used. 11. After imaging, remove the cartridge from the microscope and unload the AutoGrid in a process mirroring that of sample loading. 3.4 Loading of AutoGrids onto CryoTEM

1. The autoloader cassette is used to transfer the AutoGrids into and out of the Autoloader and is labeled 1 to 12 from bottom to top. Place the cassette in the transfer station with the top end facing to the right. Cool the station and the NanoCab. Transfer the AutoGrid box to the station. Use the AutoGrid forceps to insert the AutoGrids into the slots on the cassette so that the carbon side of the grid is facing to the right. 2. Once loaded, visually inspect to make sure that they are fully inserted. Use the AutoGrid forceps to press gently (to the right) against the top side of the AutoGrid. It springs back into position if seated properly. 3. Attach the NanoCab to the transfer station. Grasp the cassette by holding down the button on the cassette arm. Slide the cassette into the NanoCab, release the button, and withdraw the arm. 4. Gently pull the pin on the NanoCab up to make sure it is not stuck. Slide the NanoCab into the microscope slot with the cryo warning label facing out. The green “Dock” button on the left most side will light up. Press it to start the loading process. When the process is complete, take out the NanoCab and close the microscope door. 5. Wait until the temperature is ready (below 165
C). In the TEM User Interface, click “Inventory” from the Autoloader flap-out panel. After the inventory operation is complete, select a grid slot and click “Load.”

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The tuning procedures (see Note 4) described in steps 1 to 7 below are usually performed both at the beginning of the data collection session and just before starting the batch data collection described in Subheading 3.16, while steps 8 and 9 can be performed just before the batch data collection. 1. Some tuning procedures should be performed over a thin area of carbon, whereas the others should be performed over the vacuum area. To find these areas on the grid, first make sure the objective aperture is retracted (see Note 5). In SerialEM, take an 82 whole grid montage as described Subheading 3.7 below. To set the vacuum position, add a point on the hole of a broken square and label it as “vacuum” in the “Note” text box. Move to the point, turn on the low dose mode and take a “View” image to make sure it is truly over the vacuum, that is, there should be no feature in the field of view. To set the carbon position, add a point on a square, which has thinner ice, take a “View” image, move the point to a thin area of carbon, and retake an image to make sure the whole imaging area is over the carbon. Label this point as “carbon.” 2. To set the eucentric height, activate the “Record” imaging mode in SerialEM as described in Subheading 3.6. Move to the “carbon” point and find eucentricity by clicking “Eucentric-Both” from the “Tasks” menu, which will first find the rough eucentricity followed by finding the fine eucentricity. Press the Eucentric focus button on the right microscope control panel (RCP). To set the true focus, click “Set Target” from the “Focus/Tune” menu, enter a defocus value of 0 μm, and click “OK.” Click “Autofocus” from the same menu. Verify that it is truly in focus on DigitalMicrograph. Turn on “View,” and select “Live/FFT” from the “Process” menu to show the real-time fast Fourier transform (FFT) of the image. Use the Focus button on the RCP to change the defocus to a negative value such as 2 μm. Thon rings will appear on FFT. Adjust the focus back to around 0 until the rings disappear. Press the R2 button on the RCP to reset the defocus to 0. 3. Beam tilt pivotal points are focus-dependent; therefore, the alignment should be done at the eucentric focus on the “carbon” area. Lower the fluorescent screen, select “Beam tilt pp X” in the “Direct Alignments” panel on the “Tune” tab, and use the Multifunction X knob on the left microscope control panel (LCP) to superimpose the two beam spots (only appear if the alignment is initially bad) and minimize the beam movement. Click “Done.” Select “Beam tilt pp Y” and repeat the process. Select “Beam shift” and use the Multifunction X and Y knobs to center the beam to the K3 camera position (GIF entry

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aperture, which is indicated with the green circle). The green circle can be turned on or off by clicking the “GIF” symbol above the display window. 4. The objective stigmatism can be automatically corrected using Sherpa on the microscope controlling computer. While on the “carbon” area, set the defocus to 1 μm using the Focus knob. In the “Auto Functions” window of Sherpa, click “Correct” in the “Objective stigmatism” field of the “Controls” panel. The default setting is to bring the remaining astigmatism below 5 nm. Sherpa will iterate until this threshold is satisfied. 5. Coma-free alignment can also be automatically performed using Sherpa. While on the “carbon” area, set the defocus to 3 μm using the Focus knob. Click “Correct” in the “Coma” field. The default threshold for coma is 160 nm. If the procedure fails, increase the threshold to get closer, and then change it back to 160 nm to repeat the correction. After the coma-free alignment, do another round of objective stigmatism correction at the defocus of 1 μm because coma correction often introduces additional astigmatism. 6. To insert the objective aperture, lower the fluorescent screen and get out of the energy-filtered transmission electron microscopy (EFTEM) mode by pressing “EFTEM” on the “Filter” panel. Press the Diffraction button on the RCP to switch to diffraction mode. Click “Natural” below the display and use the mouse wheel to change brightness/contrast of the fluorescent screen viewer until you see the diffraction rings. On the “Apertures” panel, select the 100 μm objective aperture and click “Objective” to insert it. Click “Adjust,” use the Multifunction X and Y knobs to center the aperture around the central spot, and click “Adjust” again to finish. Press the Diffraction button again to return to imaging mode. Click “EFTEM” on the “Filter” panel to return to the EFTEM mode. Reinsert the K3 camera from the “Camera” menu in DigitalMicrograph. 7. The following three steps are for the K3 camera and should be performed over vacuum and on the “Record” imaging mode in SerialEM. The first step is to center the energy filter on the zero-loss peak (ZLP). In SerialEM, move to the “vacuum” point and stay on the “Record” mode. In DigitalMicrograph, select “Power User Mode” on the “Help” menu and the “Linear” mode on the K3 camera. Lower the fluorescent screen and change the spot size to 2 (L3 on the LCP) to increase the beam intensity. Select “Beam shift” and use the Multifunction X and Y knobs to center the beam so that it is covering the GIF entry aperture shown in the TEM User Interface display window. Lift the screen. Unblank the beam and center the energy filter by clicking “Center ZLP” on the “Tune GIF” panel.

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8. GIF needs to be tuned with a strong beam. To make sure the beam intensity is sufficient, click “View” in DigitalMicrograph and switch “Aperture” to “Mask” in the Filter Control panel. The mask should be strongly contrasting (bright hole on a dark background). Switch “Aperture” back to “Imaging” and stop “View.” Click “Tune GIF.” A “Tune Imaging Filter” dialog will pop up. Click the wheel-like symbol to open the settings, check all steps for “Full Tune” and click “OK.” Click “Full Tune” to start. The steps will run in the following order: “Image centering,” “Spectrum focus” (isochomaticity, rough tune), “Achromaticity,” “XY Magnification” (first round), “Spectrum aberrations” (isochomaticity, fine tune), “Image distortion,” and “XY Magnification” (second round). Monitor the progress in the log window during the tuning procedure to determine which steps have been completed and whether the results are acceptable. A “Tuning complete” window will pop up if it is completed successfully. If not, uncheck the completed steps and run “Full Tune” again with the remaining steps. Sometimes the “Image distortion” tuning will be worse, and a dialog will pop up with this message “Tuned distortion appears to be worse. Undo tune?” Click “Undo.” Select “XY Magnification” in the “Quick Tune” menu and run “Quick tune” to complete the tuning. 9. Gain reference is collected for the linear mode first. In DigitalMicrograph, select “Prepare Gain Reference” from the “Camera” menu. A window will pop up with the message “Collect reference images for Linear mode?”. Click “Collect Gain Reference” to proceed. After a 10 s dark image is automatically collected, adjust the beam intensity using the intensity knob on the LCP until the average count in the image is around 1280 as directed by the program. Click “Done.” After 20 images have been acquired and averaged, a new window will pop up with the message “Collect reference images for Counted mode?”. Do not click “Collect Gain Reference” yet. Lower the fluorescent screen and change the spot size to 6. Lift the screen, unblank the beam if necessary, and press “Collect Gain Reference.” After a 10 s dark image is automatically collected, a “Gain Reference Exposure Setup” window will pop up. Use the default dose rate of 15 e/pix/s (see Note 6) and a total of 6000 electrons, and click “OK.” Use the intensity knob to adjust the beam intensity until the average dose rate is around 15 e/pix/s and click “Done.” After 20 images have been acquired and averaged, a window will pop up with the message “The gain reference image has been successfully acquired and saved.” Click “OK.” The gain reference files are saved in the “Reference Images” folder in C:\ProgramData\Gatan. The most important one is the counted mode gain reference, which has the extension “.x1.m1.dm4.” Copy this file to the specified grid folder.

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1. SerialEM is usually installed on the K3 camera controlling computer (K3PC). Start the SerialEM server file FEI-SEMserver.exe on the microscope controlling computer, and then open the SerialEM program from the K3PC.

3.6 Prepare Imaging Settings in SerialEM

2. To minimize unnecessary beam exposure for dose-sensitive samples, SerialEM implements two imaging modes: the regular mode and the low dose mode. The regular mode has one exposure area and is mainly used for general tasks such as beam alignment and map acquisition. The low dose mode is mainly used for data acquisition and usually uses three exposure areas to complete the task: record, view, and focus/trial areas. The record area is where the target is located and the tilt series are collected, while the view area is centered on the record area but covers a larger area at medium magnification. The focus/ trial area is usually set to be a few μm away from the record area along the tilt axis and is used for focusing and tracking operations so that the record area is only exposed for recording the tilt images. 3. The regular mode imaging conditions can be set and saved in the Imaging States dialog as described in Steps 4 to 7 below. This dialog is used to store and return to a set of imaging conditions including both the microscope settings and imaging parameters. Table 1 listed the representative regular mode imaging states that will be used in the following sections. The magnification and dose rate are two major imaging and data collection parameters that need to be carefully considered. See Note 7 for proper choices of these two parameters. 4. To create these states for the first time, click “Open” from the “Navigator” menu to open the Navigator window (see Note 8), and then click “Open Imaging States.” The Imaging States dialog will pop up.

Table 1 Representative regular mode imaging states used in SerialEM data acquisition

Imaging Spot Illumination State # Magnification size area (μm)

Dose rate (e/pix/s)

Exposure time (s)

Slit width Binning (eV)

Defocus (μm)

1

82

9

~920

18–20

0.4

1

None

50

2

470

9

~200

18–20

0.4

1

None

50

3

4800

7

~20

18–20

0.4

1

50

40

4

15,000

7

6.3

~40

0.2

1

20

40

5

33,000

7

2.4

~40

0.1

1

20

Variable

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5. To set the microscope settings for a new imaging state, go to a “vacuum” point that is set as described in step 1 of Subheading 3.5, and then adjust the magnification (Magnification knob on the RCP), spot size (L3 button on the LCP and R3 button on the RCP), and illumination area (Intensity knob on the LCP) to obtain the desired dose rate. Be sure to lower the fluorescent screen before changing the spot size to protect the camera. Take a “Preview” or “Record” image from the Camera & Script panel to check the dose rate, which is shown on the up-left corner of the main display. The energy filter can be inserted or retracted, and the slit width can be adjusted in the Filter Control panel. 6. The imaging parameters should be set up in the “Record” tab of the Camera Parameters dialog. Open this dialog from the “Camera” menu, select the “Record” tab, enter the exposure time for the corresponding state in Table 1, and set binning to 1. In addition, set “Parameters for Acquisition” to “Single Image,” “Processing” to “Gain Normalized,” and “Operating mode” to “counting.” Make sure “Dose Fractionation mode” is unchecked and leave other fields as default. 7. Click “Add Current State” to add the current microscope and “Record” parameters as a new imaging state in the panel. Note that in this version of SerialEM (3.8.6), the defocus value is not defined in this panel but can be set separately before taking images with any of the states. An imaging state can be activated by double clicking on it or by highlighting it and clicking the “Imaging” button in the same panel. 8. Four frequently used low dose imaging modes need to be set up in the Low Dose Control panel: “View” (V) at medium magnification of 4800, and “Focus” (F), “Trial” (T), and “Record” (R) at high magnification that is used for data collection. 9. To initially set up the “View” mode, activate Imaging State 3 (4800) in the Imaging States dialog, check “Low Dose Mode” in the Low Dose Control panel, take a “View” image from the Camera & Script panel, check “Continuous update,” adjust any settings as necessary, and then uncheck “Continuous update.” 10. While still in the “View” mode, click “Copy current area mag & beam to” “R,” take a “Preview” or “Record” image to go to the record area, check “Continuous update,” and adjust the settings to be the same as Imaging State 5 (33,000) (see Note 9) as described in Steps 5 and 6 above. Select “Keep Focus and Trial identical,” click “Copy current area mag & beam to” “F” and “T,” and then uncheck “Continuous update” to finish the setup for the “Record,” “Focus,” and “Trial” modes.

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Table 2 Camera parameter setup for the low dose mode in SerialEM Dose fractionation mode

Exposure time (s)

Binning

Gain Counting normalized

Off

0.4

2

Single image

Gain Counting normalized

Off

0.4

2

Trial

Single image

Gain Counting normalized

Off

0.4

2

Record

Single image

Gain Counting normalized

Off or on

Variable

1 or 0.5

Preview

Single image

Gain Counting normalized

Off

0.1

4

Imaging mode

Parameters for acquisition

View

Single image

Focus

Processing

Operating mode

11. The imaging parameters for the above four low dose modes should be defined in the Camera Parameters dialog. In each tab, define the parameters for the corresponding low dose mode as shown in Table 2 while leaving other parameters as default. The exposure time can be variable. In the “Record” tab, “Dose Fractionation mode” can be turned on when setting up the tilt series data collection later. 12. The low dose “Record” imaging state at each magnification can also be added to the Imaging States dialog by pressing “Add Current State,” and conveniently activated later when needed. All the regular and low dose mode imaging settings generated above can be saved into individual user’s settings file and retrieved in future sessions, both from the “Settings” menu. 3.7 Collection of a Whole Grid Montage Map at Low Magnification

1. A low magnification montage map for the entire grid should first be acquired to assess the overall quality of the grid (see Note 10). Activate Imaging State 1 (82) in the Imaging States dialog and make sure a Navigator window is opened. 2. From the “Navigator” menu, open the Montage Setup dialog by clicking “Montage & Grids/Setup Full Montage.” Set the number of pieces in X to 4 and Y to 6. Check “Move stage instead of shifting image” and “Ask about making map after each montage” while leaving other fields as default. Clicking “OK” will open the “File Properties” dialog. Use the default settings and click “OK.” Save the montage map as “FullMontage82X.”

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3. Press “Start” in the Montage Controls panel. When finished, answer “Yes” to the prompt to save the map, which will appear in the item list box of the Navigator window. Save the Navigator file as well. 4. Examine the map by dragging with the left mouse button and zooming with the middle button. A very dark region would suggest that the ice there is too thick. Select those squares that appear lighter and have sharp edges, and place a point in the middle of each selected square by clicking “Add Points” in the Navigator window and then clicking on the map. These points will be automatically saved in the item list box. 3.8 Grid Square Exploration to Identify Regions of Interest

1. To further evaluate the squares for ice thickness and target presence, montage maps for individual squares are subsequently collected at 470. Click “Close” from the “File” menu to clear the 82 Montage map in the buffer. 2. In the Navigator window, click the first selected point from the 82 montage map, check “Collapse” to collapse these points into a group. Check “Acquire (A)” will mark all the points in this group with “A”. Uncheck “Collapse.” 3. Activate Imaging State 2 (470) and set the defocus to 50 μm using the Focus knob on the RCP. Click “Montage Setup” from the “File” menu, set the number of pieces in X to 2 and Y to 2, check “Move stage instead of shifting image” while leaving other fields as default. Click “OK,” and save the map as “SquareMontage470X.” 4. Click “Acquire at Items” from the “Navigator” menu, check “Rough eucentricity” and “Acquire map image or montage,” and then click “GO.” The acquired maps will appear in the item list box of the Navigator window. 5. Examine the squares by loading the map into the display through double clicking on each map item with the left mouse button. Select regions of interest by clicking “Add Polygon” and placing multiple points on the map to draw a polygon. Avoid regions that have many empty holes (appeared very bright on map), heavy contamination (ice crystals or aggregated materials), or large cell clumps. Edge regions should also be avoided because the beam can potentially hit the grid bar at high tilts. Ideally for bacterial samples, select regions that have single cells uniformly distributed with good spacing.

3.9 Set Image Shift Offset Between 4703 and the “View” Magnification (48003)

1. Image features usually will shift significantly when going between 470 and the next higher magnification to be used: the “View” magnification at 4800. To correct the shift, load a 470 map, find a featured contaminant piece, which can be

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either an ice crystal or aggregated material, add a point on the feature, label it as “Offset” on the “Note” text box, and click “Go To XYZ.” 2. When the stage movement stops, turn on “Low Dose Mode” from the Low Dose Control panel, and take a “View” image to find the feature. Note that there is a 180
rotation along the Z-axis on the images between the two magnifications. If the feature does not appear in the current view, use the right mouse button to drag the image and click “View” to examine the surrounding regions until the feature can be found. 3. With the “Offset” point selected in the Navigator window, click “Move Item” and then click on the feature to move the point to it. Click “Stop Moving” and then “Go To XYZ” to bring the “Offset” point to the center. 4. Go back to the original 470 map by loading it in the Navigator window. Clicking on the “Offset” point (orange cross) with the left mouse button will place a marker (green cross) on the same position, that is, the orange cross and green cross will overlap. 5. Select the “Offset” point in the Navigator window, click “Move Item” and then click the feature on the 470 map. This point should now be on the feature. Click “Stop Moving.” 6. With the “Offset” point still selected, click “Shift to Marker” from the “Navigator” menu and answer “Yes” to the prompt. To verify that the offset has been properly set, click “Go To XYZ” for the “Offset” point and then take a “View” image. The feature should now be in the center with the point marked on it. 3.10 Target Region Exploration at Medium Magnification

1. The regions of interest can be imaged at medium magnification to allow more detailed view on the cells and better assessment on ice thickness; thus, the next step is to acquire 4800 polygon montage maps for the regions selected from the 470 maps acquired in Subheading 3.8. Click “Close” from the “File” menu to clear the 470 montage map in the buffer. 2. To make the montage maps clearer, it may be better to more accurately set the defocus. Activate Imaging State 3 (4800), add a point on the carbon in one of the polygon regions on a 470 map, and click “Go To XYZ.” Click “Set Target” from the “Focus/Tune” menu, and enter the defocus value of 40 μm. Click “OK” and then “Autofocus” from the same menu. 3. In the Navigator window, mark “Acquire” and “New file at item” for the polygon items. A “Properties of File to Open” dialog will pop up. Check “Montaged images,” “Fit montage to polygon” and “Skip montage setup dialog when fitting in

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future.” The “Montage Setup” dialog will pop up. Check “Move stage instead of shifting image” while leaving other fields as default and click “OK” to save the file as “PolygonMontage4800X.” 4. Click “Acquire at Items” from the “Navigator” menu, check “Fine eucentricity” and “Acquire map image or montage,” and then click “GO.” The acquired maps will be saved in the item list box. 5. If the cells are more sparsely distributed, then acquiring regular montage maps at 4800 may be a better choice as this is faster. See Note 11 for the procedures. Examine the 4800 montage maps and add points or draw polygons (depending on what type of maps to acquire next in Subheading 3.11 below) on good regions. A good region is usually indicated by monodispersed cells and by holes that contain thin ice. Holes with good ice thickness often exhibit a gradient that is lighter in the center and darker around the edge. 3.11 Target Identification at High Magnification

1. To more precisely identify the targets for data collection, it is often necessary to collect maps at high magnification (15,000, 19,500 or 33,000 are frequently used). Before map acquisition, the image shift between the “View” (4800) and “Record” (15,000, 19,500 or 33,000, which is determined by the imaging state activated) magnifications need to be corrected. The procedure is essentially the same as described in Subheading 3.9 above but replace the 470 map with the 4800 map and the “View” operation with “Preview” or “Record.” Note that there is no change in image orientation along the Z-axis between these two magnifications. It may also be better to more accurately set the defocus as described in step 2 of Subheading 3.10 above but at high magnification. Set the defocus to a value between –20 to 40 μm. 2. There are three ways to obtain the high magnification maps: (1) acquiring projection images, which are suitable for smaller areas; (2) regular montaging, which is suitable for mediumsized areas; and (3) polygon montaging, which is suitable for larger areas. Which way to use depends on the concentration and complexity of the targets. 3. To acquire projection images, mark “Acquire” for the picked points on the 4800 maps. Click “Close” from the “File” menu to clear the 4800 montage map in the buffer and then “Open New” from the same menu. Save the map as “ProjectionImg33000X.” Activate Imaging State 5 (or others depending on what magnification is more suitable for the sample), click “Acquire at Items” from the “Navigator”

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menu, check “Acquire map image or montage,” and then click “GO.” 4. To acquire polygon or regular montage maps, follow the procedures described in Subheading 3.10 or the Notes section (see Note 11), but use the polygons or points picked on the 4800 maps and use high magnification (such as Imaging States 4 or 5) for imaging. To be faster, do not check “Fine eucentricity” since it has already been done at 4800 above. 5. Examine the high magnification maps and add points on the targets. Targets are identified based on their features, such as an appendage or protrusion on the cell membrane. Try to pick targets located in the holes for thinner ice. Make sure the points are not spaced too closely to prevent double exposure during data collection. 3.12 Determine the Electron Dose and Exposure Time for Data Collection

1. Before setting up the batch data collection, it is necessary to determine the proper total electron dose and calculate the exposure time for each image. Cellular samples can usually ˚ 2. Note that the tolerate a total dose of up to 100 to 200 e/A unit for the measured dose rate given by SerialEM is e/pix/s, ˚ 2/s using the pixel size (A˚). It is which can be converted to e/A easier to understand the dose calculation using this formula: Total dose (e/A˚2) ¼ [dose rate (e/pix/s) / (pixel size (A˚))2]  exposure time per image (s)  number of images. Therefore, the exposure time per image (s) ¼ [total dose (e/A˚2)  (pixel size (A˚))2] / [dose rate (e/pix/s)  number ˚ 2, the of images]. As an example, if the total dose is 163 e/A ˚ pixel size is 2.65 A at 33,000, the dose rate is 40 e/pix/s, and there are 41 images per tilt series, the exposure time ˚ 2  2.65  2.65 A˚2) / (40 e/ per image would be (163 e/A pix/s  41) ¼ 0.7 s. 2. To determine how much total dose the specimen can tolerate, collect a tilt series using a dose on the higher range and inspect the images at high tilt angles to see if there is any severe radiation damage to the sample. Severe radiation damage usually appears as bubbles in the specimen at higher tilt angles. Reduce the dose until no damage can be visualized.

3.13 Set Record Camera Parameters and File Options for Data Collection

1. The imaging parameters and file saving options are defined in the “Record” tab of the Camera Parameters dialog. Open this dialog from the “Camera” menu and select the “Record” tab. Set “Binning” to either 1 or 0.5 (superresolution mode), exposure time to 0.7 s following the example in step 1 of Subheading 3.12 above. Turn on the “Dose Fractionation mode,” and enter a “Frame time” of 0.1 s to get seven frames per tilt image.

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2. Check “Align frames” and “Save frames.” By default, the dosefractionated frames will be aligned with the “framealign” module in SerialEMCCD plugin to DigitalMicrograph, and saved as a compressed TIFF frame stack. These stacks will be further processed using the “framewatcher” program in IMOD as described in step 1 of Subheading 3.17 below. 3. Click “Set File Options,” set the file type as TIFF with LZW compression, and enter a desired base file name. If you plan to use the “framewatcher” and “alignframes” programs in IMOD for preprocessing the frames as described in step 1 of Subheading 3.17 below, check “Save unnormalized frames even if Gain Normalized is selected” to save unnormalized frames. Otherwise, if you plan to use another independent program such as MotionCor2 [54] for frame alignment, it may be more convenient to save gain normalized frames by unchecking this option. Leave other fields as default and click “OK.” 4. Click “Set Folder” to have the images saved to a “Frames” subfolder in the specified grid folder. Click “OK” to finish the setup. 3.14 Set the Tilting Scheme and Other Data Collection Parameters

1. The tilting scheme and other data collection parameters are defined in the Tilt Series Setup dialog. Select the first target point in the Navigator window and check “Tilt series.” A “Properties of File to Open” will pop up. Select “Single frame images” and click “OK.” In the next pop-up window, save the tilt series as “TS001” in the specified grid folder. If the question “There is an imaging state currently set. Do you want to restore the prior state before setting up the tilt series?” pops up, answer “No” and the Tilt Series Setup dialog will open. 2. In the “Tilt Angle Specifications” field, set “Tilt to” 60 and “End at” 60 with a base increment of either 2 or 3 degrees. A smaller range of tilt angles, such as 45 ̊ or 36 ̊ may be desirable for high-abundance targets in order to achieve higher electron dose and higher image contrast in each tilt images and thus higher resolution [55, 56]. Check “Run series in two directions from 0,” and use either the bidirectional tilting scheme or the dose-symmetric scheme. Also check “Use View for anchor.” 3. In the “Autofocus Control” field, enter a “Defocus target” between 4 andto 8 μm for cellular targets. In the “Initial and Final Actions” field, uncheck “Align to image now in Buffer A” and “Refine eucentricity.” Leave other fields as default and close this dialog. 4. Continue to mark “Tilt series” for the remaining target points.

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3.15 Define the Focus Position and Set Periodic Energy Filter Centering

1. The focus position for each target point needs to be defined individually and should be located in an area on either the left or the right side of the target point along the tilt axis, which is approximately horizontal. Load the high and medium magnification maps containing the target points and determine which side along the tilt axis can be used as the focus/trial area for each point. Avoid those areas that have heavy contamination or are close to a crack or the grid bar. 2. With the first point selected, check “Focus” in the “Define position of area” field of the Low Dose Control panel, and enter a value between 3 and 5 μm for the “Position on tilt.” The value should be negative for a focus position on the right side of the target point and positive for the left side. 3. Go through the points one by one from up to bottom and click the “Set: Focus Pos” tab in the Navigator window whenever there is a change between negative and positive values. A letter “P” will be marked on the point when the “Set: Focus Pos” tab is clicked. 4. Energy filter centering (Center ZLP) should be performed periodically at least every 2 h or every six tilt series during batch data collection. First, go to the “vacuum” point on the grid, pick multiple points inside the hole, and label them with “ZLP” in the “Note” text box. Mark “Tilt series” for these points. Move the “ZLP” points up by clicking and dragging them with the left mouse button so that there is one “ZLP” point between every six tilt series points. When SerialEM moves to these points, Center ZLP will be performed using a custom script (see Note 12).

3.16 Start the Batch Tilt Series Acquisition

1. To start the batch tilt series acquisition, highlight the first target point and click “Acquire at Items” from the “Navigator” menu. In the “Initial Actions after Moving Stage” field, check “Realign to item” and “Fine eucentricity.” Also check “Run script” here and select the custom Center ZLP script from the drop-down menu. In the “Primary Task” field, check “Acquire tilt series.” Click “GO” to start. 2. Raw frames from each exposure/tilt angle are stacked into a compressed TIFF file and saved, along with a command file with the extension .pcm and a metadata file with the extension . mdoc, into the “Frames” subdirectory created in step 4 of Subheading 3.13 above. In the meantime, the entire tilt series are stacked and saved as a .mrc file, along with the .pcm and . mdoc files, into the specified grid folder.

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The raw images output by SerialEM are gain normalized, motion corrected, and stacked into a tilt series for tomogram reconstruction (see Note 13) as follows. 1. Install Cygwin and IMOD on the K3PC according this guide: h t t p s : // b i o 3 d . c o l o r a d o . e d u / i m o d / d o c / g u i d e . html#SettingUpWindows. Open a Cygwin terminal, go to the “Frames” subdirectory containing the raw data and run the command: framewatcher -gpu 0 -bin 1 -po 1024 -pr rawTIFF -thumb alignedJPG -dtotal 0 (see Note 14). 2. The aligned frames from each tilt angle are stacked into a single MRC file (*ali.mrc) and saved in the “Frames” subdirectory, whereas the raw data files will be moved to the “rawTIFF” subdirectory. The program will scan the “Frames” subdirectory for newly appeared eligible files every 5 s and can be terminated by pressing “Ctrl” and “C” on the keyboard. 3. The aligned images are then processed by the Caltech Tomography Database and Automatic Processing Pipeline (see Note 13), which can back up the raw data, reconstruct the tomograms and present them on a web browser via the internet. All these tasks are executed by running a submission script db_proc.slurm. To begin with, mount the data drive of the K3PC onto the data processing workstation. Create a subdirectory “Pipeline” in the grid folder. Move the tilt series stacks (. mrc) and the associated .pcm and .mdoc files described in step 2 of Subheading 3.16 to the “Pipeline” subdirectory. 4. On the data processing workstation, edit the required parameters and folder options in the db_proc.slurm script, and execute it with the command sbatch db_proc.slurm. 5. The script will monitor the appearance of finished tilt series stacks in the “Pipeline” subdirectory, copy the aligned frame stacks (along with the raw frames in the “rawTIFF” subdirectory) in the “Frames” subdirectory to the database, create tilt series stacks (images binned by 2 if acquired in superresolution mode) from the aligned frame stacks, reconstruct the tomograms, and generate movies and thumbnail images of the central slice from the tomograms. A binning factor of 4 (or 8 for data collected at the superresolution mode) is typically applied through the script to reduce file size and allow for faster processing. 6. The data can be accessed remotely via a web browser with password protection. The movies can be checked at the beginning and periodically throughout the data collection session to allow for any collection parameters adjustments. Checks should include whether or not the defocus is optimal, the target of

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interest is within the field of view, and for any other artifacts as a result of suboptimal collection parameters. 3.18 Manual Tomogram Reconstruction

1. Usually, the tilt series need to be manually processed to generate an optimal reconstruction for further analysis and subtomogram averaging. The major procedures for reconstructing the tomograms using Etomo in IMOD is described here. A detailed description of the processes and tutorials can be found here: https://bio3d.colorado.edu/imod/doc/ tomoguide.html. 2. Start the program with the command etomo and select “Build Tomogram”. Input the tilt series generated by the automatic pipeline in step 5 of Subheading 3.17 above, click “Scan Header,” and enter the “Fiducial diameter” of 10 nm. Click “Create Com Scripts” to start the reconstruction workflow. If at any point Etomo is prematurely closed, it can be restarted and continued from the last unfinished step by running the command etomo *edf. 3. The “Pre-processing” step removes X-rays and other artifacts that are extreme low or high values. The default parameters should suffice for most cases. Click “Find X-Rays,” “Create Fixed Stack,” and then “Use Fixed Stack.” 4. The “Coarse Alignment” step uses cross-correlation to coarsely align the images of the tilt series which facilitates automatic tracking of gold fiducials in the next step. Click “Calculate Cross-Correlation” and then “Generate Coarse Aligned Stack.” The alignment can be evaluated by clicking “View Aligned Stack in 3dmod.” For a poorly aligned stack, defining a sub-area of the images for alignment in the advanced panel can often yield an improvement. 5. The “Fiducial Model Gen.” step generates a seed model of gold fiducials and tracks them across the images in the tilt series. In the “Seed Model” tab, select “Make seed and track” and “Generate seed model automatically.” Check “Refine Center with Sobel filter.” Set “Sobel sigma relative to bead size” to 0.12 and “Overall low-pass filter cutoff (/nm)” to 0.35. Enter a total number of seed points between 10 and 30, and click “Generate Seed Model.” The model can be examined by clicking “Open Seed Model.” Manually delete undesirable points by selecting them and pressing the “Delete” key, and add more points around the targeted structure by clicking on the gold beads using the middle mouse button. 6. In the “Track Beads” Tab, check “Refine center with Sobel filter,” “Fill seed model gaps,” and “Local tracking.” Set “Sobel sigma relative to bead size” to 0.12, “Overall low-pass filter cutoff (/nm)” to 0.35, and “Local area size” to between

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100 and 1000. Click “Track Seed Model” to start the tracking, and then “Fix Fiducial Model” to fix poorly tracked beads manually. 7. The “Fine Alignment” step applies the rotation, translation, and scaling (magnification) needed to transform the images for better alignment. Select “Do not sort fiducials into two surfaces for analysis” and leave other setting as default. Check “Compute Alignment.” The program will track the beads and compare its tracked trajectory with its theoretical trajectory. View the log window for the residual error mean and standard deviation. Typically, good tracking yields values less than 1 and 0.5 for the mean and standard deviation, respectively. However, tracking with values larger than these may still be acceptable. 8. Inappropriately placed fiducials can be viewed by clicking “View/Edit Fiducial Model” and corrected manually. Save the model by pressing the “S” key and compute the alignment again. Repeat until all fiducials are well placed and the residual error is acceptable. 9. The “Tomogram Positioning” step allows to repositioning the tomogram to make it as flat and as thin as possible. Although repositioning the tomogram to generate a thin sample saves space, this would enforce interpolation, which may degrade the reconstruction quality. In addition, saving space is usually not an issue; therefore, it is unnecessary to adjust the tomogram position in most cases. For small bacterial cells, a value of 600 for “Positioning tomogram thickness” is usually sufficient to encompass the sample. Click “Create Final Alignment” to finish. 10. The “Final Aligned Stack” step generates the aligned stack. Set “Aligned image stack binning” to 4 for enhanced contrast and click “Create Full Aligned Stack.” Generally, creating the full aligned stack is sufficient. However, if higher resolution subtomogram averaging is needed, it may be helpful to correct the CTF and/or apply a 2D filter. If the target structure is proximal to gold beads, which might interfere with downstream processes, it will be helpful to erase the gold beads. 11. The “Tomogram Generation” step can reconstruct the tomogram using either Weighted Back Projection or Simultaneous Iterative Reconstruction Technique (SIRT). A weighted back projection will usually suffice. Select “Back Projection” and set the “Tomogram thickness in Z” to 600. Select to apply a SIRTlike filter to increase contrast (typically 5–15 iterations). Highfrequency filtering removes higher-resolution information for increased contrast and signal-to-noise ratio. Select “Standard

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Gaussian” and use the default parameters. Click “Generate Tomogram” to complete. 12. In the “Post-processing” step, the “Volume Range” can be defined to crop out a part of the tomogram containing the target(s) and reduce the file size. Otherwise, simply uncheck “Convert to bytes” and check “Rotate around X-axis.” Click “Trim Volume” to complete. 3.19 Particle Picking Using IMOD

1. The 3D subvolumes of the macromolecules, usually referred to as particles, are picked and extracted by IMOD here. Open the tomogram in the IMOD slicer window from the terminal using the command: 3dmod -S tomogram.rec. 2. Click the checkerboard button in the top left of the slicer window to interpolate pixels, click the “keep current image or model point centered” square, and set the slicer thickness to 10. Activate the “model” mode in the small 3dmod window. 3. Go to “Edit” and select “Angles” to open the window of saved angles. Go to “Edit” and select “Object” and then “Type.” In the newly opened window, ensure that “scattered” is selected. 4. Identify the T4PM by identifying the location where a pilus contacts the outer membrane of the bacterial cell (Fig. 2a, b). Left click on the point where the pilus contacts the outer membrane to center the red crosshair over this point. Rotate the Z-axis until the pilus becomes vertical above the outer membrane. Next, rotate the X-axis until the outer and inner membranes are clear and at their closest point to each other (Fig. 2b, c). Middle click on the outer membrane to place a model point. 5. Click “new” at the top of the Slicer window. This will save a new angle in the Angles window. If you click “save” you will

Fig. 2 Picking and manual alignment of T4PM particles. (a) Identification of pili on the bacterial cell (surrounded by red dashed box). (b) Reorientation of the T4PM particle. Identify where the pilus contacts the outer membrane (white arrow), center the red cross on the outer membrane at that location, and rotate along the Z-axis to make the outer membrane horizontal. (c) Manual orientation of the T4PM particle for subsequent subtomogram averaging. Rotate along the X-axis until the outer and inner membrane are at their closest distance to each other. (Images were adapted from [11])

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overwrite the last saved angle. Once this has been done for all the pili in the tomogram, click “File” and select “Save model as.” Save this as a .mod file. 6. Repeat steps 1–5 for all the desired tomograms. 7. Next, particle coordinates and angles must be extracted for importing into Dynamo. To extract coordinates, we convert all the model files to text files in the terminal using the command model2point in.mod out.txt. 8. To extract particle angles, we use a custom shell script that uses the command imodinfo to extract Slicer angles from the model files and creates a motive list with the command slicer2MOTL, which converts X-Y-Z slicer angles into Z(1)-Z(3)-X (2) Euler angles and writes them into columns 17–19, respectively, of the motive list. 3.20 Subtomogram Averaging Using Dynamo

Dynamo is used to perform subtomogram averaging as follows (see Note 15). 1. Open MatLab from the terminal with the command matlab. In MatLab, navigate to the Dynamo installation directory and type the command dynamo_activate to activate Dynamo for use in MatLab. 2. In MatLab, navigate to the directory to be used for the subtomogram averaging project. We typically do this inside a directory called “Dynamo.” 3. Initialize the Dynamo project by creating a “catalog.” This can be done with the command dcm -create catalogName, where “catalogName” is the name of your Dynamo project. We typically give this the name of the structure we are resolving, such as T4PM. 4. Next, tomograms from which particles were picked must be imported into the Dynamo catalog. This can either be done in the Dynamo Catalog Manager (dcm) GUI by typing the command dcm and selecting the “Catalogue” tab and clicking “Browse for new volume” or in the MatLab command line with the command dcm -c catalogName -at /path/to/tomogram.rec, where “/path/to/tomogram.rec” is the full path to your tomogram file. 5. In addition to importing tomograms into the catalog, you must create a “table map” file with the extension .doc that contains a list of the paths to each tomogram in the same order in which they were imported into the catalog (see Note 16). 6. Next, particle coordinates must be imported into Dynamo from IMOD and cropped from the tomograms in the Dynamo catalog. To do this, we use a custom MatLab script, which, in

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Table 3 Euler angle conversions between PEET and Dynamo PEET

Dynamo

Z(1), column 17

-Z’ (column 9)

Z(3), column 18

-Z (column 7)

X(2), column 19

-X (column 8)

brief, reads the model point coordinates from the text files output by model2point into a MatLab matrix, creates a Dynamo model of type “General” with these coordinates, and crops these points from their respective tomograms into a specified particles folder with an associated metadata table named “crop.tbl” (see Note 17). For this project, the particle box size is 90 nm (corresponding to 116 pixels with a pixel size of 7.8 A˚) 7. To import particle angles, we use a custom MatLab script that imports the motive lists that contain the particle angles and converts the PEET-convention angles (Z[1], Z[3], X[2] from columns 17–19, respectively) into Dynamo-convention angles (referred to as Z, X, Z’ or drot, dtilt, and narot). It then assigns these angles to the particles from the respective tomograms in the “crop.tbl” file (columns 7, 8, and 9, respectively) (Table 3). 8. A subtomogram alignment project can begin once the tomograms, particle coordinates, and angles are all imported into the Dynamo project. To perform this, initialize the GUI with the command dcp. Enter the desired name of alignment in the box and hit enter (we usually begin with “run1” and successively increase the number with additional alignments). 9. One by one input the information for each tab as they activate: Particles: browse and select the particle directory. Table: browse and select the “crop.tbl” file inside the particle directory. Template: browse and select the zero-search average or a desirable particle. Mask: first click “Use default masks” at the bottom. Then under alignment mask either select the mask editor to create an ellipsoid mask or browse and select a custom mask. Numerical parameters: fill in the information as shown in Table 4. This will iteratively align all the particles and average the best 2=3 of particles for use as a new template for the next iteration. The final iteration will average all

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Table 4 Numerical parameters for run1 and run1_sym2 Parameter

run1

run1_sym2

Iterations

3

1

References

1

1

Cone aperture

12

3

Cone sampling

2

0.75

Azymuth rotation range

12

3

Azymuth rotation sampling

2

0.75

Refine

4

3

Refine factor

2

2

High pass

2

2

Low

42

42

Symmetry

c1

c1

Particle dimensions

116

116

Shift limits

333

111

Shift limiting way

1

2

Separation in tomogram

0

0

Basic MRA

0

0

Threshold parameter

1

1

Threshold modus

5

5

particles with the new angles and translations from the previous iteration. Computing environment: select “GPU under MatLab” if you have enabled the GPU. At the bottom, enter the number of CPUs for parallel processing in the “Parallel processing” box if the parallel computing toolbox is enabled in MatLab. Check: click this to allow Dynamo to check for inconsistencies in the project parameters. Unfold: click this to create a MatLab script, necessary to perform the subtomogram alignment and averaging. Run: click this to execute the subtomogram alignment project. 10. Settings and results from each subtomogram alignment project have a specific directory structure within that of your project, for example, “run1” in this case. All results can be found in “run1/results.” Within this is a directory for the results of each iteration. The final average and metadata table (referred to as

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the “refined table”) will be in the directory “run1/results/ ite_0003/averages/.” Every average is named according to the number of the template to which it was aligned as well as the iteration. For example, in this project, the final average will be called “average_ref_001_ite_0003.em.” Similarly, the final refined table will be called “refined_table_ref_001_ite_0003. tbl” (see Note 18). 11. For the T4PM, twofold symmetry can be observed throughout the subtomogram average and thus we will apply twofold symmetry. To do this, we use a custom MatLab script, which, in brief, in the MatLab workspace duplicates each particle, rotates the duplicated particles by 360/n degrees (where n in this case is 2) along the Z’ Euler angle (column 9 from the refined table), and re-crops them from the tomograms into a new particle directory. 12. After creating symmetrized particles, create a new subtomogram alignment project in the dcp called “run1_sym2.” Fill in the parameters as in step 9 above. Here, the particles and table will be those from the directory containing symmetrized particles. Furthermore, the template will be the average from run1. For the numerical parameters, fill in the information as shown in Table 4. 13. Upon completion of this alignment project, the resulting average will be saved as “average_ref_001_ite_0001.em” within “run1_sym2/results/ite_0001/averages/.” 3.21 Resolution Estimation and Structural Analysis

1. Resolution estimation in Dynamo can be performed using “Adaptive bandpass filtering” (see Note 19). First, initialize the dcp with your alignment project with the command, in this example, dcp run1_sym2. 2. Go to the “Multireference” tab, hover over “Adaptive filtering (“gold standard”)” and select “Derive a project.” This will create a new alignment project with the same name + _eo. In this case, it will be called “run1__sym2_eo.” In the dcp GUI, type “run1_sym2_eo” into the “Project” box and hit enter to begin editing parameters for the adaptive bandpass filtering project. 3. Go to the “Multireference” tab, hover over “Adaptive filtering (“gold standard”) and select “Edit parameters for Adaptive Filtering run.” Enter the following information for each parameter: Adaptive filtering: 1 Adaptive bandpass threshold: Set to 0.143 for gold standard Fourier shell correlation resolution estimation. Recompute rotated reference: 1

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Adaptive initial lowpass: Set to the Fourier pixels that correspond to your estimation of resolution. In this case with an estimation of ~4 nm, set to 23 (see Note 20). Adaptive bandpass pushback: 0 Symmetrized resolution: 1 Adaptive bandpass even/odd: 0 4. In an adaptive bandpass project, all input files are preloaded from the project from which this was derived. Therefore, it is only necessary to edit the “numerical parameters.” Input the same parameters as the alignment project (Table 4). Change the following parameters: Iterations: set to a number between 5 and 10 References: 2 Threshold parameter: 1 Threshold modus: 5 5. With these parameters modified, the adaptive bandpass filtering project can now be executed. Click “check,” then “unfold,” and finally “run.” 6. As each iteration completes, the resulting Fourier shell correlation at 0.143 (estimated resolution) will be output into a text file called “bandpass_resolution.txt” within the results directory of the project folder (in this case, “run1_sym2_eo/ results/bandpass_resolution.txt”). The results are in Fourier pixels. 7. To show the resulting plot, read all the fsc files that are generated by the project into the MatLab workspace with the command fsc = dread(‘run1_sym2_eo/results/ite*/averages/ bandpass*.fsc’). This command reads in the fsc curve, which is an array of values, for each iteration into the variable “fsc.” 8. Next, plot each fsc curve with the command dfsc_show({fsc}, ‘style’, 1, ‘apix’, 7.8). This plots the FSC curve with ang˚. stroms on the X-axis and a pixel spacing of 7.8 A 9. When analyzing a structure in which various conformations are resolved or mutant structures are resolved, it is beneficial to visualize the structural differences. To do this, we can create a 3D difference map between the wild-type and mutant structures in ChimeraX. Prior to creating 3D difference maps in ChimeraX, it is necessary to align the structures of interest, invert the contrast of the voxels within the volumes, convert them to .mrc file format, and normalize their densities. The first three will be performed in Dynamo, and the density normalization will be done using an IMOD command. 10. To align the volumes in Dynamo, one volume will be the “particle” and one will be the “template.” Read both volumes

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Table 5 Abbreviations and descriptions for subtomogram alignment parameters Parameter abbreviation

Parameter

Description

cr

Cone range

Angular range to search for the Z (drot) and X (dtilt) Euler angles

cs

Cone sampling

Angular step size to perform during the cone search

ir

In plane (azymuth) Angular range to search for the Z’ (narot) Euler angle range

ir

In plane (azymuth) Angular step size to perform during the azymuth search step

dim

Box dimensions

The particle size for alignment. Can be reduced to speed up the alignments.

rf

Refine factor

Number of times to reduce the angular search range for refinement

rff

Refine parameter

The factor by which the angular search range is reduced at each refinement level Range[i + 1] ¼ sampling  rff[i] Sampling[i + 1] ¼ Sampling[i] / rff

lim

Translational search limits

Pixels to search in X, Y, and Z

limm

Search mode

The way in which the center of the particle is defined prior to the translational search

into the MatLab workspace with the command particle = dread(‘volume1.em’); and template = dread(‘volume2. em’). In this case, “volume1.em” and “volume2.em” are the paths to the files containing the wild-type and mutant structures. 11. Align the volumes with the command oal = dynamo_align (particle, template, ‘cr’, 20, ‘cs’, 5, ‘ir’, 20, ‘is’, 5, ‘dim’, 116 ‘rf’, 5, ‘rff’, 2, ‘lim’, [10 10 10], ‘limm’, 1). See Table 5 for descriptions of the subtomogram alignment parameters. This command will perform an angular search 20 degrees for all Euler angles with a step size of 5 and reduce by half for five iterations. It will also perform a translational search of 10 pixels in each dimension. The resulting aligned volumes will be stored in the variable “oal.aligned_particle” and “oal. aligned_template.” 12. Invert the contrast of the aligned particle volumes using the commands volume1 = -oal.aligned_particle”; “volume2 = -oal.aligned_template. This will invert the volumes such that protein density appears white. Save them to .mrc files with the

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command dwrite(volume1, ‘T4PM_wt.mrc’); dwrite(volume2, ‘T4PM_mutant.mrc’). 13. In the terminal, use the command densmatch T4PM_wt.mrc T4PM_mutant.mrc T4PM_mutant_normalized.mrc to normalize the mutant T4PM average to the wild-type T4PM average. 14. Open UCSF ChimeraX. Load in the volumes for the wild-type T4PM and the normalized, mutant T4PM subtomogram averages. 15. Type the command volume subtract #1 #2, where #2 is the identifier for the mutant average and #1 is the identifier for the wild-type average. The resulting map will be the mutant structure subtracted from the wild type structure, revealing densities that are missing from the mutant structure.

4

Notes 1. The cells need to be mixed with gold fiducials prior to vitrification so that the tilt images can be more precisely aligned by tracking the gold fiducials during tomogram reconstruction. Before usage, the gold fiducials need to be coated with BSA to prevent aggregation. 2. The carbon film surface of the grids is hydrophobic and generally will expel an aqueous solution. A glow discharge treatment with air will make the carbon film surface negatively charged and thus more hydrophilic. This treatment is usually performed within 1 h of sample application. 3. The horizontal and vertical positions of the grid for blotting should first be set using an empty grid according to the manufacturer’s manual. 4. Proper alignment of the microscope and preparation for the camera is critical for obtaining high quality data [57–59]. The protocol assumes that the microscope operates in Nanoprobe EFTEM mode at SA mode magnifications and that the gun alignment, C2 aperture centering, and C2 condenser stigmatism correction have been performed on a monthly basis. Most procedures should be performed using the imaging mode preset with the planned data acquisition conditions, for example, magnification, spot size, and illumination area. Here, SerialEM is used for data collection; therefore, its “Record” mode (described in step 10 of Subheading 3.6) will be used. 5. The “Objective” button in the “Apertures” panel of the “EFTEM” tab in the TEM User Interface is yellow when the aperture is inserted or gray when retracted. If it is in the inserted state, click the yellow “Objective” button to retract it.

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6. If the actual dose on the camera post-specimen will be significantly different from 15 e/pix/s, take the gain reference for the counted mode at the actual post-specimen dose. A gain reference taken at the wrong dose rate may lead to camera artifacts in the corrected images. 7. Low and medium magnifications are used to assess the grid quality and identify regions of interest on the squares and a dose rate of 18–20 e/pix/s over vacuum is appropriate. High magnifications are used for target picking and data acquisition and an average count of 15 e/pix/s post-specimen is optimal for the K3 camera. For cellular samples, a dose rate of ~40 e/ pix/s over vacuum is approximately good to achieve the electron count of ~15 e/pix/s post-specimen. Regarding the choice of magnification for data acquisition, the theoretical maximum resolution at a given magnification is twice the pixel size (the Nyquist frequency). In practice, a pixel size smaller than the theoretical value is recommended [58]. For example, if the target resolution for the sample is 1 nm, then the pixel size should be 5 A˚ theoretically or smaller in practice. We usually use 33,000 magnification (pixel size ~2.65 A˚) for imaging bacterial samples. 8. The Navigator window is the “Data Management Center,” which is used to store and track stage positions and maps. It is also used for a variety of image acquisition-related operations for the stored items. SerialEM occasionally crashes; therefore, it is important to frequently save the Navigator items to a file. The saved Navigator file can be retrieved by “Read & Open” from the main menu. 9. 33,000 is the most frequently used magnification for tilt series acquisition for bacterial surface nanomachines in our laboratory. If higher magnification such as 64,000 or 81,000 is needed, the imaging settings can be adjusted as described in steps 5 and 6 above and updated with “Continuous update.” 10. A grid needs to be imaged at incremental magnifications in a step-wise fashion to acquire maps which can be used for evaluating its quality and properly identifying the targets of interest for tilt series acquisition. 11. To acquire regular montage maps at 4800, add points on the center of the target regions on the 470 maps and mark “Acquire” in the Navigator window. Activate Imaging State 3 (4800), and set the defocus to 40 μm as described in step 2 of Subheading 3.10 or manually by using the Focus knob on the RCP. Click “Montage Setup” from the “File” menu, set the number of pieces in X to 3 and Y to 3, check “Move stage instead of shifting image” while leaving other fields as default.

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Click “OK,” and save the file as “RegularMontage4800X.” Click “Acquire at Items” from the “Navigator” menu, check “Fine eucentricity” and “Acquire map image or montage,” and then click “GO.” 12. The scripts in this chapter are available for sharing upon request. 13. We usually use the “framewatcher” program in IMOD and the Caltech Tomography Database and Automatic Processing Pipeline [60] to perform real-time data processing, storage, and tomogram reconstruction for the collected tilt series. Tomograms can also be reconstructed directly by IMOD or other software packages. “Framewatcher” is a Python program in IMOD that can watch for either the command files (.pcm) or the frame stacks (. tif) appearing in a directory and automatically run the “alignframes” program to align and sum the raw frames in batch. 14. Commands are written in bold text. Italicized parts of commands are variable and correspond to the proper file or parameter name. 15. A variety of software packages are currently available to perform subtomogram averaging, such as PEET, Dynamo, PyTom, emClarity, EMAN2, and Relion [52, 61–65]. Furthermore, workflows have been established to perform near atomic resolution subtomogram averaging, such as the WARP/M/ Relion pipeline [66]. For cellular structures, we generally use Dynamo to perform subtomogram averaging. More automated workflows in Dynamo have been established for subtomogram averaging of certain structures [67, 68]. As a prerequisite to this protocol, MatLab must be installed to run Dynamo. Alternatively, Dynamo can be run in a stand-alone mode (not discussed here). Documents for the usage of Dynamo can be found here: https://wiki.dynamo. biozentrum.unibas.ch/w/index.php/Main_Page. We recommend doing the Advanced Starters Guide to familiarize yourself with the Dynamo and MatLab interfaces prior to beginning your own project: https://wiki.dynamo. biozentrum.unibas.ch/w/index.php/Advanced_starters_ guide. 16. More information about the “table map” file can be found here: https://wiki.dynamo.biozentrum.unibas.ch/w/ index.php/Tomogram-table_map_file. 17. To create a general type model and crop points in Dynamo, the following commands are used (the first two commands are used in a for loop that iterates through every IMOD model file):

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m = dmodels.general(); %Creates a model of type General in the variable ‘m’. m.addPoint(coordinate); %Where ‘coordinate’ is an array containing X, Y, and Z values for one model point. m.updateCrop(); dcm(‘-c’, catalog, ‘-i’, volNum, ‘-am’, m, ‘-modelname’, modelName); % where “catalog” is the name of the catalog, “volNum” is the number of the volume to, which this model belongs, and “modelName” is the model to be saved in Dynamo. t = m.grepTable(); % Creates a particle table for the model points and stores in variable “t.” dtcrop(‘tomogram.rec’, t, ‘particleDirectory’, 116); % where “tomogram.rec” is the path to the tomogram file from which this model is cropped, “particleDirectory” is the name of a directory into which particles will be cropped, and “116” is the size of the particle to crop. 18. As iterations of an alignment project are completed, the results will be populated within the project directory. For example, if we assign four iterations to “run1,” we can check the results of each iteration as they are completed. In IMOD, results can be opened from “run1/results/ite_000X/averages/average_ref_001_ite_000X.em,” where “X” is the iteration number. As alignments are running, we generally check the results from intermediate iterations if we are testing new numerical parameters to ensure that the particles are aligning well. 19. More information about “Adaptive bandpass filtering” can be found here: https://wiki.dynamo.biozentrum.unibas.ch/w/ index.php/Adaptive_bandpass_filtering. 19. Resolutions in Dynamo are generally represented in the units of Fourier pixels (fp). To convert to resolution in A˚, use the formula: ResolutionðÅÞ ¼ ½box size ðpixÞ  pixel size ðÅÞ=fp

Acknowledgments We thank Dr. Shrawan Kumar Mageswaran and William D. Chen for technical assistance during the development of the workflow and other members of the Chang lab for helpful discussions. This work is supported in part by a David and Lucile Packard Fellowship for Science and Engineering (2019-69645), a Pennsylvania Department of Health FY19 Health Research Formula Fund, a Mark Foundation For Cancer Research ASPIRE Award

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(19-044-ASP), and a NIH R01 (1R01GM134020-01A1) to Y.-W. C.; a Taiwan National Science and Technology Council (NSTC) grant (110-2926-I-001-501) to E.-M.L., M.-C.H., Y.-W.C.; a NSTC postdoc fellowship (110-2811-B-001-524) to Y.-W.C.; a NIH R35 grant (R35-GM130302) to B.E.B.; a NIH F30 fellowship (F30-CA261198) to K.K.; and a Martin and Pamela Winter Infectious Disease Fellowship to M.M. This work was performed in part at the Beckman Center for Cryo-Electron Microscopy at the University of Pennsylvania, which is supported by the Arnold and Mabel Beckman Foundation. References 1. Persat A, Inclan YF, Engel JN et al (2015) Type IV pili mechanochemically regulate virulence factors in Pseudomonas aeruginosa. Proc Natl Acad Sci 112(24):7563–7568. https://doi. org/10.1073/pnas.1502025112 2. Craig L, Forest KT, Maier B (2019) Type IV pili: dynamics, biophysics and functional consequences. Nat Rev Microbiol 17(7): 429–440. https://doi.org/10.1038/s41579019-0195-4 3. Nakamura S, Minamino T (2019) Flagelladriven motility of bacteria. Biomolecules 9(7): 279 4. Wang J, Brodmann M, Basler M (2019) Assembly and subcellular localization of bacterial type vi secretion systems. Annu Rev Microbiol 73(1):621–638. https://doi.org/10. 1146/annurev-micro-020518-115420 5. Costa TRD, Harb L, Khara P et al (2021) Type IV secretion systems: advances in structure, function, and activation. Mol Microbiol 115(3):436–452. https://doi.org/10.1111/ mmi.14670 6. Ellison CK, Whitfield GB, Brun YV (2021) Type IV pili: dynamic bacterial nanomachines. FEMS Microbiol Rev. https://doi.org/10. 1093/femsre/fuab053 7. Miyata M, Robinson RC, Uyeda TQP et al (2020) Tree of motility – a proposed history of motility systems in the tree of life. Genes Cells 25(1):6–21. https://doi.org/10.1111/ gtc.12737 8. Chang Y-W, Shaffer CL, Rettberg LA et al (2018) In vivo structures of the Helicobacter pylori cag type IV secretion system. Cell Rep 23(3):673–681. https://doi.org/10.1016/j. celrep.2018.03.085 9. Ge P, Scholl D, Prokhorov NS et al (2020) Action of a minimal contractile bactericidal nanomachine. Nature 580(7805):658–662.

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Chapter 19 Twitching Motility Assays of Lysobacter enzymogenes OH11 Under a Light Microscope Bingxin Wang, Xiaolong Shao, and Guoliang Qian Abstract Bacterial twitching motility is a peculiar way of adherence and surface translocation on moist solid or semisolid surfaces. Although the twitching motility has been detected in various flagellated bacteria, such as Pseudomonas aeruginosa, it has been rarely detected in flagella-less bacteria like Lysobacter enzymogenes, a natural predator of filamentous fungi. Here, by using a strain OH11 of L. enzymogenes as a model system, we describe a convenient method for observing the twitching motility, with fewer steps and better repetition than conventional methods. This new method provides important technical support for the motile study of Lysobacter. Key words Lysobacter enzymogenes, Twitching motility, Type IV pili, Surface translocation

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Introduction Bacterial motility is one of the main characteristics of bacteria, and twitching motility is a particular surface translocation mode that occurs in flagellated and non-flagellated bacteria [1]. The twitching motility depends on retractable type IV pili (T4P) but not on the flagellum, by which many motile bacteria use to swim in viscous liquids and move on solid surfaces, and is important for host colonization and other forms of colonization behavior [2, 3]. Consisting of a helical arrangement of pilin proteins encoded by the pilA gene, T4P is a hair-like protein appendage approximately 5–8 nm in diameter and extending to several micrometers in length [3]. Most of them are located at one or both poles of the cell and are regulated by a variety of factors. The T4P is also involved in bacterial adhesion and colonization on host cells, biofilm formation, horizontal gene transfer, and so on [4, 5]. Twitching assays have been widely used in flagellated Pseudomonas aeruginosa with two slightly modified methods [6].

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Lysobacter enzymogenes OH11 (OH11), a member of the Xanthomonadaceae family, is a natural predator of filamentous fungi. Due to the lack of flagellar genes, OH11 is incapable of displaying flagellar-driven motility but can assemble T4P for twitching motility. Our recent findings have shown that OH11 still remains the flagellar type III secretion system (FT3SS) on its genome [7]. To investigate the function of the FT3SS in OH11, we delete the FT3SS ATPase gene fliI from the OH11 genome and show that the fliI gene is required for the secretion of antifungal toxins and the major pilus subunit, PilA, indicating that the FT3SS can positively regulate the twitching motility [7, 8]. Thus, we believe that twitching is the main motility manner of OH11 to facilitate its colonization behavior. However, a simple and reproducible method for detecting the twitching motility of Lysobacter is still lacking to date. In this chapter, we describe a visualized method to detect the twitching motility of Lysobacter enzymogenes. We use a thin layer of solidified nutrient medium on a glass coverslip as the twitching motility substrate and observe the results under optical microscopy.

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Materials All solutions were prepared using ultrapure water (by purifying deionized water to achieve a conductivity of 18 M Ω/cm at 25 ° C) and analytical grade reagents, and all reagents were prepared and stored at room temperature (unless otherwise noted). The test strains are wild type of Lysobacter enzymogenes OH11 and a derivative mutated for fliI. 1. 0.15% (w/v) Tryptic Soy Broth agar: 1.5 g/L BD Tryptic Soy Broth, 16 g/L agar. 2. 3% (w/v) Tryptic Soy Broth: 3 g/L BD Tryptic Soy Broth. 3. Luria-Bertani broth: 10 g/L Tryptone, 5 g/L yeast extract, 10 g/L NaCl. 4. Luria-Bertani agar: 10 g/L Tryptone, 5 g/L yeast extract, 10 g/L NaCl, 16 g/L agar. 5. Yellow plastic disposable pipette tips, sterile. 6. Blue plastic disposable pipette tips, sterile. 7. Fixed volume pipettes. 8. Plastic disposable sterile inoculation loop, 10 μL. 9. Petri plates: 90 mm in diameter, sterile. 10. Autoclave bottle, minimum volume 300 mL. 11. Autoclave bottle, minimum volume 100 mL. 12. Microwave oven.

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13. Coverslips (20 × 20 mm). 14. Microscope slides (75 × 25 mm) (see Note 1). 15. Filter paper: 80 mm in diameter. 16. Metal flat mouth tweezers. 17. Metal scalpel. 18. Heating incubators. 19. 15 mL disposable plastic tubes, sterile. 20. Incubator shaker.

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Methods Carry out all procedures at room temperature unless otherwise mentioned. All sterilized items should be cooled to room temperature before use. A flow chart is shown in Fig. 1.

3.1 Activation of Culture

1. Streak each test strain onto an LB agar plate and incubate them at 28 °C for 2 days. 2. Inoculate a single colony of each Lysobacter enzymogenes strain grown on the LB agar plate into a 20 mL volume of autoclaved LB medium and grow them at 28 °C for 16 h with shaking at 220 rpm. 3. Pipette 200 μL of the cell culture to 20 mL of autoclaved Tryptic Soy Broth medium and grow them at 28 °C with shaking at 220 rpm until the cell density reaches an OD600 of approximately 1.0.

3.2 Preparation of Solid Medium

1. Place coverslips, microscope slides, filter paper, flat mouth tweezers, and scalpel in a lunch box, and autoclave at 121 °C for 20 min (15 psi should be sufficient). 2. Melt 100 mL of autoclaved 0.15% (w/v) Tryptic Soy Broth agar using a microwave (see Note 2). 3. Dispense 10 mL of molten Tryptic Soy Broth agar into a petri plate using a sterile 15 mL disposable centrifuge tube to make solid medium. Gently swirl it to spread evenly and make sure it is level (see Note 3). 4. Leave the agar to set for 30 min. 5. Using an autoclaved scalpel, cut the agar into rectangles of about 22 × 40 mm.

3.3 Microscope Slide Preparation

1. Place the cut medium on a microscope slide with flat-mouth tweezers, and make sure that it is level (see Note 3). 2. Place a piece of autoclaved filter paper in a new petri plate using autoclaved tweezers.

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Fig. 1 Schematic diagram of the experimental procedure of twitching motility assays of Lysobacter enzymogenes

3. Pipette 2 mL of autoclaved water into each plate to fully soak the filter paper. 4. Place the microscope slide with medium on the soaked filter paper using autoclaved tweezers. 5. Using autoclaved flat mouth tweezers, gently touch one edge of the coverslip to the surface of the inoculum prepared in Subheading 3.1. 6. Slowly place the coverslip onto the agar to minimize the bubble formation between the coverslip and the agar.

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Fig. 2 Microscopic twitching assay of Lysobacter enzymogenes. Mobile cells at the colony margin are observed on the wild-type strain OH11 plate, but are not observed on the ΔfliI plate. Scale bars: 10 μm

7. Cover the petri dish and mark the lid. 8. Incubate them at 28 °C for at least 24 h to observe clear cell traces in microscopic analysis. 3.4 Microscope Observation

1. Place a slide on the microscope stage. 2. Find the edge of solid medium touched to the inoculum with 10× low magnification dry objective lens. 3. Switch to 63× oil immersion objective lens and focus. 4. Move the slide very slowly to find the area of twitching of Lysobacter enzymogenes. We select the wild-type strain OH11 and ΔfliI as test strains. As shown in Fig. 2, the twitching of ΔfliI compromises compared with the wild-type strain OH11.

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Notes 1. For high-resolution images, we recommend to use clean and high-transmittance coverslips and glass slides. 2. It is important to visually monitor the solution to make sure that it is not boiling. Make sure the solution is completely thawed before use. 3. It is important to use a spirit level to ensure that the surface is flat, or this can make it very difficult to take good pictures by a camera if the surface of the slide or agar is uneven.

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Acknowledgments This work was supported by the Natural Key Research and Development Program (2022YFD1400200 to G.Q.), the National Natural Science Foundation of China (U22A20486, 32072470 and 31872016 to G.Q.), Science and technology project of Shanxi Branch of China National Tobacco Corporation (KJ-2022-04 to G.Q.). This work was also supported in part by JSPS KAKENHI (JP19H03182, JP22H02573, and JP22K19274 to T.M.). References 1. Henrichsen J (1972) Bacterial surface translocation: a survey and a classification. Bacteriol Rev 36(4):478–503. https://doi.org/10.1128/br. 36.4.478-503.1972 2. Henrichsen J (1983) Twitching motility. Annu Rev Microbiol 37:81–93. https://doi.org/10. 1146/annurev.mi.37.100183.000501 3. Mattick JS (2002) Type IV pili and twitching motility. Annu Rev Microbiol 56:289–314. https://doi.org/10.1146/annurev.micro.56. 012302.160938 4. Craig L, Pique ME, Tainer JA (2004) Type IV pilus structure and bacterial pathogenicity. Nat Rev Microbiol 2(5):363–378. https://doi.org/ 10.1038/nrmicro885 5. Nudleman E, Kaiser D (2004) Pulling together with type IV pili. J Mol Microbiol Biotechnol 7(1–2):52–62. https://doi.org/10.1159/ 000077869

6. Turnbull L, Whitchurch CB (2014) Motility assay: twitching motility. Methods Mol Biol 1149:73–86. https://doi.org/10.1007/978-14939-0473-0_9 7. Fulano AM, Shen D, Kinoshita M et al (2020) The homologous components of flagellar type III protein apparatus have acquired a novel function to control twitching motility in a non-flagellated biocontrol bacterium. Biomol Ther 10(5). https://doi.org/10.3390/ biom10050733 8. Fulano AM, Shen D, Zhang EH et al (2020) Functional divergence of flagellar type III secretion system: a case study in a non-flagellated, predatory bacterium. Comput Struct Biotechnol J 18:3368–3376. https://doi.org/10.1016/j. csbj.2020.10.029

Chapter 20 Live Cell Imaging of the Twitching Motility of Cyanobacteria by High-Resolution Microscopy Daisuke Nakane Abstract Many cyanobacteria show directional movement either toward or away from light sources. The cell movement, also known as twitching motility, is usually driven by type IV pili (T4P), a bacterial molecular machine. The machine generates a propulsion force through repeated cycles of extension and retraction of pilus filaments. Here, I describe a phototaxis assay for observing Synechocystis sp. PCC6803 and Thermosynechococcus vulcanus at the single-cell level with optical microscopy. By adding fluorescent beads, I also describe a method how to visualize the asymmetric activation of T4P during phototaxis. Key words Type IV pili, Phototaxis, Motility, Optical Microscopy, Signal transduction

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Introduction Type IV pili (T4P) are ubiquitous molecular machines in bacteria that drive twitching motility, protein secretion, and DNA uptake [1]. Twitching motility refers to cell movement over surfaces such as glass or agar that is propelled by repeated cycles of the extension and retraction of pilus filaments (Fig. 1a) [2–4]. T4P dynamics are powered by two ATPases for the assembly and disassembly of pilin and are regulated by a signal transduction system that responds to a variety of environmental stimuli [5]. Many cyanobacteria use light as an energy source via photosynthesis, and optimal light conditions are crucial for efficient photosynthesis. The cells move over surfaces in a T4P-dependent manner, either into their preferred light habitat or away from harmful or stressful environments, with the movement directional along the optical axis of the light stimulus [6–8]. Recent study has shown that cyanobacteria recognize the light direction via their cell body as a micro-sized optical lens to sense the light intensity difference along the optical axis [9–12]. This intensity difference

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Fig. 1 T4P-dependent phototaxis in cyanobacteria. (a) Schematic of twitching motility. A cell moves by repeated cycles of extension, attachment, and retraction of T4P filaments. (b) Diagram of the experimental setup. Green and blue LEDs are simultaneously applied through dichroic mirrors into the left side of the cells on the microscope stage as a lateral light stimulus for phototaxis. The position of the cells is visualized by near-infrared light. (c) Upper: Bright-field image of Synechocystis sp. PCC6803 and their moving trajectories for 240 s on a glass substrate coated with collodion. Black arrows indicate the direction of the lateral blue light for phototaxis. Lower: A kymograph of the cell movements along the optical axis of the lateral illumination is presented. The directional movements of the cells are indicated over time by the tilted lines. The tilted lines from the left-upper to the right-lower side present negative phototaxis

induces directional cell motility by the activation of T4P machinery with an asymmetric manner along the optical axis of the light stimuli [9, 10, 12]. In cyanobacteria like Synechocystis sp. PCC6803 and Thermosynechococcus vulcanus, the asymmetric T4P activation is visualized by a nanosized fluorescent sulfate bead as a probe: T4P filaments at the front side of the directional movement capture the beads and retract them toward the cell surface (Fig. 2a) [10, 12]. Since the diameter of single pilus is only 7 nm and it is difficult to detect its dynamics with optical microscopy [3, 13, 14], the bead assay can be a useful method that is applicable to various cyanobacteria. In this chapter, we describe a method for observing cell behaviors in Synechocystis sp. PCC6803 and T. vulcanus with optical microscopy (Fig. 1b, c). We also describe a bead assay for visualizing the extension and retraction of the T4P filament (Fig. 2b–d). These assays can be useful for observing T4P-dependent cell motility in response to light illumination on a short time scale.

2

Materials Prepare all solutions using ultrapure water and analytical grade reagents. Sterilize the solutions (autoclave for 20 min at 121 °C) prior to use and store all reagents at 4 °C (unless indicated otherwise).

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Fig. 2 Visualization of T4P dynamics through fluorescent beads in Synechocystis sp. PCC6803. (a) Schematic of the bead assay. (b) Typical trajectories of beads in a series of images. The time is indicated at the bottom left of the image. A bead moves toward the immobilized cell located on the left side. (c) Trajectories of the beads. Beads tracked over a time interval of 0.1 s are presented. The circle at the center indicates the position of the cell. (d) Time course of the distance between the bead and the center of the cell. The velocity was measured based on the time course of the displacement 2.1 Bacterial Strains and Growth Medium

1. Synechocystis sp. PCC6803: PCC-P strain (see Note 1). 2. Thermosynechococcus vulcanus: Positive phototactic strain of T. vulcanus [12] (see Note 2). 3. Solution I for BG11: 0.30 g ferric ammonium citrate and 0.05 g Na2 EDTA 2H2O in 100 mL of water. 4. Solution II for BG11: 30 g NaNO3, 0.78 g K2HPO4, and 0.73 g MgSO4 in 1 L of water. 5. Solution III for BG11: 1.43 g CaCl2 in 100 mL of water. 6. Solution IV for BG11: 2.00 g Na2CO3 in 100 mL of water. 7. Solution A6 for BG11: 2.86 g H3BO4, 1.81 g MnCl2 4H2O, 0.22 g ZnSO4 7H2O, 0.08 g CuSO4 5H2O, 0.021 g Na2MoO4 2H2O, and 0.05 g Co(NO3)2 6H2O in 1 L of water. 8. TES buffer for BG11: 1 M TES-KOH, pH 7.5. 9. BG11 liquid medium [12]: 2 mL of Solution I, 50 mL of Solution II, 2 mL of Solution III, 1 mL of Solution IV, 1 mL of Solution A6, and 20 mL of 1 M of the TES buffer to 1 L of water. 10. Incubator at 30 °C and 45 °C. 11. Flask: 100 mL. 12. Silicon plug. 13. Fluorescent lamp to grow cells. 14. Shaker: 180–200 rpm. 15. Powermeter for measuring the light intensity. 16. Spectrophotometer, 750 nm.

2.2 Glass Chamber Assembly

1. Coverslip: 22 × 40 mm. 2. Coverslip: 18 × 18 mm.

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3. Double-sided tape. 4. Nail polish. 5. Collodion: 2% (w/v) in isoamyl acetate. 2.3 Optical Microscopy

1. Inverted microscope (e.g., IX73, Olympus). 2. 10×, 20×, and 100× Objective lens (e.g., UPLFLN10 × 2, LUCPLFLN20× and UPLSAPO100 × O2PH, Olympus). 3. CMOS camera (e.g., Zyla 4.2, Andor or DMK33U174, Imaging Source). 4. Bandpass filter (e.g., FBH850/40, Thorlabs) for applying vertical illumination. Visualize the cell by infrared light from a halogen lamp with the bandpass filter. 5. Optical filter (e.g., FITC-5050A, Semrock) for visualizing the fluorescent beads. 6. Thermoplate (e.g., TP-110R-100, Tokai Hit) for heating the microscope stage at 45 °C. 7. Blue and green light LEDs (e.g., M450LP1, M530L3, Thorlabs) with peak wavelengths at 450 and 530 nm, respectively, for lateral light stimuli. 8. Optical table. 9. Spectrometer. 10. Software for image analysis such as ImageJ (http://rsb.info. nih.gov/ij/).

2.4

Bead Assay

1. Sulfate beads: 200 nm in diameter (FluoSpheres sulfate polystyrene microspheres CatNo F8848, Thermo Fisher), containing fluorescent dye for excitation/emission at 505/515 nm. Dilute 1/100 in fresh BG11 liquid medium. 2. Filter paper.

3

Methods Carry out all procedures at room temperature unless otherwise specified.

3.1 Phototaxis on a Short Time Scale Under Optical Microscopy

1. Add 25 mL of the BG11 liquid medium to a 100 mL flask (see Note 3) [15]. 2. Inoculate 1 mL of the cells in the flask, and then seal the flask with a silicon plug. 3. Incubate the flask at 30 °C for Synechocystis sp. PCC6803 and 45 °C for T. vulcanus, then shake the flask at 180–200 rpm under moderate fluorescent light. Adjust the fluence rate of the light to 20 μmol m-2 s-1 by using a power meter (see Note 4).

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4. Measure the optical density (OD) of the culture at a wavelength of 750 nm with a spectrophotometer. Prepare the cell culture to an OD750 of 0.5–1.0 for the following experiments. 5. Assemble a glass chamber with two coverslips and two pieces of double-sided tape to act as a thin channel for observing the sample under optical microscopy (see Note 5). The height and width of the channel are approximately 100 μm and 10 mm, respectively, resulting in a 20 μL volume for the channel space. 6. Pour the cell culture into the glass chamber (see Note 6). 7. Seal both ends of the chamber with nail polish to ensure that the sample does not dry out. 8. Place the chamber on the microscope stage at room temperature (RT) for Synechocystis sp. PCC6803 and 45 °C for T. vulcanus. Set up a thermoplate to heat the chamber. 9. Visualize the cells in the sample chamber with vertical light illumination at a wavelength of 850 nm with a halogen lamp and a bandpass filter (see Note 7). Adjust the fluence rate to 1 μmol m-2 s-1 by using a power meter (see Note 8). 10. Apply LED light illumination at an angle of 5 degrees with the lateral side of the microscope stage. Collimate the LED light with a condenser lens. Adjust the blue light to have a fluence rate of approximately 1000 μmol m-2 s-1 to induce negative phototaxis in Synechocystis sp. PCC6803 (Fig. 1c) [10] (see Note 9). Adjust the green light to have a fluence rate of approximately 70 μmol m-2 s-1 to induce positive phototaxis in T. vulcanus [12]. Adjust the green and blue light to have fluence rates of approximately 70 μmol m-2 s-1 to induce negative phototaxis in T. vulcanus [12] (see Note 10). 11. Visualize the cell with vertical light through the 10× objective lens, and record the cell image with a camera at 1 s intervals for at least 5 min. 12. Turn on the lateral light illumination to observe the cells move in response to the lateral illumination during the recording. Remove the lateral light with another bandpass filter placed between the objective lens and the tube lens so that the camera does not detect the light (see Note 11). 13. Convert the sequential images of the cells into a TIF file without compression. Analyse the data with ImageJ (http:// rsb.info.nih.gov/ij/) and its plugins, multitracker, and particle tracker (see Note 12). 3.2 Visualizing T4P Dynamics with Beads

1. Prepare the cell in the sample chamber in the same way as steps 1–6 in Subheading 3.1 (see Note 13). 2. Pour the cell culture into the glass chamber and incubate them on the microscope stage for 3 min at RT for Synechocystis sp. PCC6803 and 45 °C for T. vulcanus.

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3. Place a piece of filter paper on one side of the tunnel in the chamber and add sulfate beads solution to replace the solution in the sample chamber (see Note 14). 4. Seal both ends of the chamber with nail polish to ensure that the sample does not dry out. 5. Apply lateral light illumination at an angle of 5 degrees with the lateral side of the microscope stage in the same way as a step 10 in Subheading 3.1. Use lateral blue light to excite the fluorescent beads. 6. Visualize the green fluorescence due to the beads with an optical filter (see Note 15). 7. Record a sequential image through the 40× objective lens with a camera for 2 min at 0.1 s intervals to observe whether the beads show directional movements towards or away from the cell surface (Fig. 2b) (see Notes 16 and 17).

4

Notes 1. Synechocystis sp. PCC6803 is known to exist in several distinct substrains, and all strains were derived from the same isolate deposited in the Pasteur Culture Collection (PCC) [17]. The positive phototaxis strain used here was isolated from the original PCC strain based on the direction of positive phototactic movement on agar plates, and designated as PCC-P [16]. Specific mutations in these strains that may contribute to the altered phenotype of these strains have been reported by whole genome sequencing [18, 19]. 2. The original strain of T. vulcanus is NIES-2134 in the Microbial Culture Collection at the National Institute for Environmental Studies (NIES, https://mcc.nies.go.jp/). The complete genome has been sequenced (AP018202). Since the original strain exhibited a heterogeneous phototaxis phenotype under optimal growth conditions, a clone isolated from the original strain is used based on the direction of positive phototactic movement on agar plates under moderate light [20]. 3. Frozen stock is prepared by cell culture. The cell culture grown in BG11 liquid medium to an OD750 of 1.0, is mixed with dimethyl sulfoxide (DMSO) at a final concentration of 5% (v/v), and is then kept at -80 °C. 4. Lower light illumination is used when the cells are initially grown from frozen stock. The cell culture is reinoculated in fresh BG11 every week. 5. For Synechocystis sp. PCC6803, the coverslip is coated with collodion. The collodion is diluted to a concentration of 2%–

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0.007% (w/v] in isoamyl acetate. A 2 μL volume of the solution is coated over a cover slip and air-dried before use. The collodion coating increases cell movement in Synechocystis sp. PCC6803. 6. The Synechocystis sp. PCC6803 cells produce polysaccharides without shaking [21], which inhibit binding to the glass surface for phototaxis. The T. vulcanus cells form aggregates at low temperature [22, 23], which inhibits single-cell observations. These cell cultures should be maintained with shaking at a proper temperature. 7. A heat absorption filter is removed from the halogen lamp, and a bandpass filter is placed in the light path to apply vertical light at a wavelength of 850 nm. Near-infrared light has no significant effect on phototaxis [6, 24]. The wavelength of the light can be verified by a spectrometer if necessary. 8. The contrast of the image can be increased by an aperture stop if necessary. The refractive index of the cyanobacteria is slightly higher than that of other bacteria [25], and the cell image shows clear contrast even under bright-field microscopy. 9. Lateral red light illumination induces positive phototaxis in the cells grown on the BG11 agar plate [9], but not in cells grown in the BG11 liquid medium for Synechocystis sp. PCC6803. The lateral blue light illumination described here was used for negative phototaxis with Synechocystis sp. PCC6803. 10. Simultaneous illumination with two LEDs, blue and green, is accomplished with dichroic mirrors [12], which enable visualization of the directional switch from positive to negative phototaxis [12]. 11. Without the bandpass filter, the light scattering of the cell due to the lateral light illumination can be visualized [10, 12]. The scattering is greater for the forward side of the light direction than for the backwards side, which is responsible for the light direction recognition in the cyanobacteria and the so-called micro-optics effect [9, 10, 12, 26]. 12. The Synechocystis sp. PCC6803 phototaxis is random on a time scale of 1 min. Cell observation for at least 5 min should be required to detect directional movement (Fig. 1c). 13. There are many ways to analyze cell behavior (Fig. 1c). For example, single-cell tracking is possible with an ImageJ plugin. Kymographs can be useful for obtaining complete phototaxis observations, as they present the directional movement of cells as tilted lines over time [12]. 14. The bead assay is only applicable to sulfate beads. For carboxylate-modified beads and amine-modified beads, directional movement toward or away from the cell is not observed [10, 12].

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15. The collodion is diluted to a concentration of 2% (w/v) to 0.2% (w/v) in isoamyl acetate, and the solution was coated over a cover slip as described in Note 5. This collodion coating immobilized the Synechocystis sp. PCC6803 cells on the glass surface and effectively inhibits the nonspecific binding of fluorescent beads to the glass surface. 16. In Synechocystis sp. PCC6803, lateral blue light illumination activates T4P dynamics asymmetrically at the forward side of the light axis, resulting in negative phototaxis (Fig. 2c). 17. To calculate the velocity of T4P extension and retraction, the distance between the bead and the center of the cell is measured, and the displacement is plotted during the observation. The distance both increases and decreases linearly over a certain period of time, indicating the extension and retraction of T4P, respectively (Fig. 2d).

Acknowledgments This work was supported by KAKENHI (16H06230, 20H05543, 21K07020, 22H05066) and funds from the Noguchi Institute to DN. References 1. Korotkov KV, Sandkvist M, Hol WGJ (2012) The type II secretion system: biogenesis, molecular architecture and mechanism. Nat Rev Microbiol 10:336–351. https://doi.org/ 10.1038/nrmicro2762 2. Merz AJ, So M, Sheetz MP (2000) Pilus retraction powers bacterial twitching motility. Nature 407:98–102. https://doi.org/10. 1038/35024105 3. Skerker JM, Berg HC (2001) Direct observation of extension and retraction of type IV pili. Proc Natl Acad Sci U S A 98:6901–6904. https://doi.org/10.1073/pnas.121171698 4. Miyata M, Robinson RC, Uyeda TQP et al (2020) Tree of motility – a proposed history of motility systems in the tree of life. Genes Cells 25:6–21. https://doi.org/10.1111/gtc. 12737 5. Craig L, Forest KT, Maier B (2019) Type IV pili: dynamics, biophysics and functional consequences. Nat Rev Microbiol 17:429– 440. https://doi.org/10.1038/s41579-0190195-4 6. Choi J-S, Chung Y-H, Moon Y-J et al (1999) Photomovement of the gliding cyanobacterium Synechocystis sp. PCC 6803. Photochem

Photobiol 70:95–102. https://doi.org/10. 1111/j.1751-1097.1999.tb01954.x 7. Yoshihara S, Suzuki F, Fujita H et al (2000) Novel putative photoreceptor and regulatory genes required for the positive phototactic movement of the unicellular motile cyanobacterium Synechocystis sp. PCC 6803. Plant Cell Physiol 41:1299–1304. https://doi.org/10. 1093/pcp/pce010 8. Bhaya D, Takahashi A, Grossman AR (2001) Light regulation of type IV pilus-dependent motility by chemosensor-like elements in Synechocystis PCC6803. Proc Natl Acad Sci U S A 98:7540–7545. https://doi.org/10.1073/ pnas.131201098 9. Schuergers N, Lenn T, Kampmann R et al (2016) Cyanobacteria use micro-optics to sense light direction. elife 5:e12620. https:// doi.org/10.7554/eLife.12620 10. Nakane D, Nishizaka T (2017) Asymmetric distribution of type IV pili triggered by directional light in unicellular cyanobacteria. Proc Natl Acad Sci U S A 114:6593–6598. https://doi.org/10.1073/pnas.1702395114 11. Wilde A, Mullineaux CW (2017) Lightcontrolled motility in prokaryotes and the

Live Imaging of Twitching Motility of Cyanobacteria problem of directional light perception. FEMS Microbiol Rev 41:900–922. https://doi.org/ 10.1093/femsre/fux045 12. Nakane D, Enomoto G, B€ahre H et al (2022) Thermosynechococcus switches the direction of phototaxis by a c-di-GMP dependent process with high spatial resolution. eLife 11:e73405. https://doi.org/10.7554/eLife.73405 13. Tala` L, Fineberg A, Kukura P et al (2019) Pseudomonas aeruginosa orchestrates twitching motility by sequential control of type IV pili movements. Nat Microbiol 4:774–780. https://doi.org/10.1038/s41564-0190378-9 14. Koch MD, Fei C, Wingreen NS et al (2021) Competitive binding of independent extension and retraction motors explains the quantitative dynamics of type IV pili. Proc Natl Acad Sci U S A 118. https://doi.org/10.1073/pnas. 2014926118 15. Stanier RY, Kunisawa R, Mandel M et al (1971) Purification and properties of unicellular blue-green algae (order Chroococcales). Bacteriol Rev 35:171–205. https://doi.org/ 10.1128/br.35.2.171-205.1971 16. Yoshihara S, Geng X, Okamoto S et al (2001) Mutational analysis of genes involved in pilus structure, motility and transformation competency in the unicellular motile cyanobacterium Synechocystis sp. PCC 6803. Plant Cell Physiol 42:63–73. https://doi.org/10.1093/pcp/ pce007 17. Rippka R, Deruelles J, Waterbury JB et al (1979) Generic assignments, strain histories and properties of pure cultures of Cyanobacteria. Microbiology 111:1–61. https://doi. org/10.1099/00221287-111-1-1 18. Kanesaki Y, Shiwa Y, Tajima N et al (2012) Identification of substrain-specific mutations by massively parallel whole-genome resequencing of Synechocystis sp. PCC 6803. DNA Res 19:67–79. https://doi.org/10.1093/dnares/ dsr042 19. Trautmann D, Voß B, Wilde A et al (2012) Microevolution in cyanobacteria:

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re-sequencing a motile substrain of Synechocystis sp. PCC 6803. DNA Res 19:435–448. https://doi.org/10.1093/dnares/dss024 20. Enomoto G, Okuda Y, Ikeuchi M (2018) Tlr1612 is the major repressor of cell aggregation in the light-color-dependent c-di-GMP signaling network of Thermosynechococcus vulcanus. Sci Rep 8:5338. https://doi.org/10. 1038/s41598-018-23628-4 21. Maeda K, Okuda Y, Enomoto G et al (2021) Biosynthesis of a sulfated exopolysaccharide, synechan, and bloom formation in the model cyanobacterium Synechocystis sp. strain PCC 6803. elife 10:e66538. https://doi.org/10. 7554/eLife.66538 22. Kawano Y, Saotome T, Ochiai Y et al (2011) Cellulose accumulation and a cellulose synthase gene are responsible for cell aggregation in the cyanobacterium Thermosynechococcus vulcanus RKN. Plant Cell Physiol 52:957–966. https://doi.org/10.1093/pcp/pcr047 23. Enomoto G, Nomura R, Shimada T et al (2014) Cyanobacteriochrome SesA is a diguanylate cyclase that induces cell aggregation in Thermosynechococcus. J Biol Chem 289:24801– 24809. https://doi.org/10.1074/jbc.M114. 583674 24. Kondou Y, Nakazawa M, Higashi S-i et al (2001) Equal-quantum action spectra indicate fluence-rate–selective action of multiple photoreceptors for photomovement of the thermophilic cyanobacterium Synechococcus elongatus. Photochem Photobiol 73:90–95. https://doi. org/10.1562/0031-8655(2001) 0730090eqasif2.0.Co2 25. Aas E (1996) Refractive index of phytoplankton derived from its metabolite composition. J Plankton Res 18:2223–2249. https://doi. org/10.1093/plankt/18.12.2223 26. Yang Y, Lam V, Adomako M et al (2018) Phototaxis in a wild isolate of the cyanobacterium Synechococcus elongatus. Proc Natl Acad Sci U S A 115:E12378–E12387. https://doi.org/10. 1073/pnas.1812871115

Part V Adhesion-Based Gliding Motility of Bacteria

Chapter 21 Isolation and Visualization of Gliding Motility Machinery in Bacteroidota Satoshi Shibata and Daisuke Nakane Abstract Many members of the phylum Bacteroidota (formerly called Bacteroidetes) adhere to and move on solid surfaces. This type of bacterial motility is called gliding and does not involve the conventional bacterial motility machinery, such as flagella and pili. To understand the mechanism of gliding motility of some Bacteroidota bacteria such as a soil bacterium Flavobacterium johnsoniae and a marine bacterium Saprospira grandis, the gliding motility machines of these two bacteria have been analyzed by electron microscopy with negative staining. Here, we describe methods to directly observe the gliding motility machinery in Bacteroidota by transmission electron microscopy. Key words Transmission electron microscopy, Negative staining, Osmotically shocked cell, Gliding motility, Phylum Bacteroidota

1

Introduction Transmission electron microscopy (TEM) with negative staining is one of the fastest and easiest methods to observe purified proteins and bacterial extracellular structures and is simpler than highly developed cryo-electron microscopy, which is now becoming popular for high-resolution structural analysis for macromolecular protein complexes. However, because of the thickness of intact cell body, direct observation of in situ structures is still challenging, even for bacterial cells. Osmotic shock has been used to visualize structures located in or on the cell membrane, such as the flagellar basal body and secretion apparatus, that is, types III and IV secretion systems [1–3]. Changes in osmotic pressure cause the cells to rupture, thereby leaking cellar contents. As a result, the thickness of the cell body becomes much thinner, which becomes more transparent for TEM. This osmotic shock method can be used to visualize the gliding machinery of Bacteroidota bacteria by TEM.

Tohru Minamino et al. (eds.), Bacterial and Archaeal Motility, Methods in Molecular Biology, vol. 2646, https://doi.org/10.1007/978-1-0716-3060-0_21, © The Author(s), under exclusive license to Springer Science+Business Media, LLC, part of Springer Nature 2023

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Fig. 1 Electron micrographs of F. johnsoniae gliding SprB adhesion filaments. (a) SprB filaments approximately 150 nm long are distributed on the cell surface. Black arrowheads indicate SprB filaments extending from the cell surface. (b) Negatively stained image of SprB filaments isolated from the cell. Bars, 200 nm

Bacteroidota have the ability to adhere to solid surfaces, where cells can move back and forth; occasionally, a cell stands and pivots in its place. By combining these movements, a cell can travel across a surface as fast as approximately 5 μm/s [4]. Cells floating in solution cannot swim but only exhibit Brownian motion. This adhesion-based motility is known as gliding motility. The molecular mechanism of the Bacteroidota gliding motility is distinct from that of conventional bacterial motilities that use either flagella or pili, as well as the gliding motility of myxobacteria and Mycoplasma [5]. The gliding machinery of Bacteroidota consists of three parts: a cell surface adhesin, a helical track associated with the outer membrane, and a proton-driven motor [6, 7]. In F. johnsoniae, the adhesin is an approximately 150 nm-long SprB filament anchored to the outer membrane (Fig. 1a) [8, 9]. The SprB filament is propelled along a left-handed closed helical loop on the cell surface. The attachment of moving SprB filaments to the solid surface results in the translocation of the cell body [9]. Various sections of the helical track have been observed by TEM with negative staining, quick-freeze-replica electron microscopy, and cryo-electron tomography. Examples include bundled filaments covering the surface of S. grandis [2, 10], a crystalline ladder-like structure [11], and a patch-like structure near the base of the outer membrane in F. johnsoniae [8]. Although the components of the helical track are not clear, the lipoprotein GldJ appears to be arranged in a helical pattern within the cell envelope in F. johnsoniae [12], suggesting that GldJ may be a component of such a helical track. The components of gliding motility and type IX secretion system (T9SS) share similarities in their amino acid sequences [13]. Structures of the proton-driven gliding motor complex

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GldLM of F. johnsoniae and the T9SS secretion motor complex PorLM of the nonmotile oral pathogen Porphyromonas gingivalis exhibit structural similarities [7]. These findings suggest that both the gliding machinery and T9SS machinery may be evolutionary related, similar to the relationship between flagella and type III secretion systems [14]. Here, we describe methods for visualizing the machinery responsible for the gliding motility of Bacteroidota by TEM with negative staining. In addition, a method for isolating SprB filaments from F. johnsoniae cells is also described.

2

Materials Prepare all buffers and staining solutions using ultrapure water (prepared by purifying deionized water to attain a sensitivity of 18 MΩ-cm at 25 °C) and analytical grade reagents. Use deionized water for culture media preparation. Sterilize media and buffers by autoclaving at 121 °C for 20 min. Store all reagents at 4 °C unless otherwise indicated. Refrigerated centrifuge with fixed angle rotor is used for centrifugation.

2.1

Cell Preparation

1. F. johnsoniae ATCC17061 (see Note 1). 2. Casitone-yeast extract (CYE) plate: 1% (w/v) polypeptone, 0.5% (w/v) yeast extract, 0.1% (w/v) MgSO4.7H2O, 10 mM Tris–HCl pH 7.5, and 1.5% (w/v) agar. 3. CYE liquid medium: 1% (w/v) polypeptone, 0.5% (w/v) yeast extract, 0.1% (w/v) MgSO4•7H2O, and 10 mM Tris–HCl pH 7.5. 4. Motility medium (MM): 0.33% (w/v) polypeptone, 0.17% (w/v) yeast extract, 0.1% (w/v) MgSO4•7H2O, and 10 mM Tris–HCl pH 7.5 (see Note 2). 5. S. grandis str. Lewin (see Note 3) 6. Marine broth plate: Difco™ marine broth 2216, 0.5% (w/v) tryptone and 1.5% (w/v) agar. 7. Marine broth liquid medium: Difco™ marine broth 2216 and 0.5% (w/v) tryptone. 8. Glycerol stock solution: 30% (w/v) glycerol. 9. Shaker incubator. 10. Spectrophotometer.

2.2 Isolation of SprB Filaments from F. johnsoniae

1. Phosphate buffered saline (PBS): 137 mM NaCl, 2.68 mM KCl, 8.1 mM Na2HPO4, 1.47 mM KH2PO4, pH 7.4. 2. 10% (v/v) Triton X-100. Sterilization is not required.

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3. 10% (w/v) n-dodecyl-β-D-maltoside (dodecyl maltoside). Sterilization is not required. 4. Ammonium sulfate. 2.3 Osmotically Shocked Cells

1. Sucrose solution: 0.5 M sucrose, 0.15 M Tris–HCl, pH 7.5. 2. Ultrapure water. 3. MgCl2 solution, 50 mM. Sterilization is not required. 4. Electron microscopy grade paraformaldehyde solution (16%). 5. Electron microscopy grade glutaraldehyde solution (10%).

2.4

TEM

1. Carbon-coated copper 200 mesh electron microscopy (EM) grid (see Note 4). 2. Glow discharge equipment for EM grids. 3. UA staining solution: 1% (w/v) uranyl acetate solution (see Note 5). 4. AM staining solution: 2% (w/v) ammonium molybdate solution (see Note 5). 5. PTA staining solution: 2% (w/v) phosphotungstic acid in ultrapure water, adjusted to pH 7.0 with 1 N NaOH (see Notes 5 and 6). 6. Whatman grade 6 filter paper. 7. Transmission electron microscope operated at an accelerating voltage of 80 kV (see Note 7).

3 3.1

Methods Cell Preparation

1. Inoculate F. johnsoniae cells on a CYE plate. Incubate the plate for more than 24 h at 25 °C (see Note 8). 2. Preculture of F. johnsoniae: Inoculate a single colony from a CYE plate into 5 mL of CYE liquid medium and incubate it at 25 °C for 16 h with shaking at 200 rpm until an optical density at 600 nm (OD600) reaches approximately 2.0. 3. Cell culture for the observation and isolation of F. johnsoniae SprB filaments: Inoculate the cell culture into a fresh CYE liquid medium at a dilution of 1:100 and incubate it at 25 °C for 6 h with shaking at 200 rpm until the OD600 value reaches approximately 1.0. 4. Cell culture for generating osmotically shocked F. johnsoniae cells: Inoculate the precultured medium into MM at a dilution of 1:100 and incubate it at 25 °C for 4 h with shaking at 200 rpm until the OD600 value reaches approximately 0.5 (see Note 9).

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5. Inoculate S. grandis cells on a marine broth plate. Incubate the plate for more than 24 h at 25 °C (see Note 10). 6. Preculture of S. grandis: Inoculate a single colony from a marine broth plate into marine broth liquid medium and incubate it at 25 °C for 16 h with shaking at 100 rpm (see Note 11) until the OD600 value reaches approximately 2.0. 7. Cell culture used for generating osmotically shocked S. grandis cells: Inoculate the preculture into a fresh marine broth liquid medium at a dilution of 1:100 and incubate it at 25 °C for 6 h with shaking at 100 rpm until the OD600 value reaches approximately 1.0. 8. Frozen stock of cells: F. johnsoniae and S. grandis cells can be frozen and stored at -80 °C in liquid media containing 15% glycerol. Mix equal amounts of the glycerol stock solution with cultured cells at an OD600 of approximately 1.5. Dispense the mixture into cryo tubes and freeze them at -80 °C (see Note 12). 3.2 Observation of SprB Filaments on the Cell Surface of F. johnsoniae by TEM

1. Harvest cells from a CYE cell culture by centrifugation at 8000 g for 10 min and suspend the cell pellet in a fresh CYE liquid medium. 2. Apply the cell suspension to an EM grid and incubate the grid for 5 min at room temperature (RT). 3. Remove approximately 90% of the liquid from the grid surface by filter paper blotting from the edge of the grid (see Note 13). The following blotting procedures are performed in the same way unless indicated otherwise. 4. Apply PBS containing 3% (v/v) formaldehyde and 0.1% (v/v) glutaraldehyde to the grid and fix the cells for 10 min at RT. 5. Remove liquids by blotting with filter paper and apply PBS. 6. Repeat step 5 three times to remove the fixing solution completely. 7. Apply AM staining solution to the grid and stain the cells for 1 min at RT. 8. Remove the liquids completely by blotting with filter paper and air dry the grid. 9. Observe negatively stained samples by an electron microscope operated at 80 kV. 10. Figure 1a shows a typical example image of negative stained SprB filaments on the cell surface of F. johnsoniae (see Note 14).

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3.3 Isolation of SprB Filament from F. johnsoniae

1. Harvest cells from a 200 mL CYE cell culture by centrifugation at 8000 g for 10 min. 2. Resuspend cells in 40 mL of PBS. 3. The following steps should be conducted at 4 °C unless otherwise indicated. 4. Add 10% (v/v) Triton X-100 to the suspension at a final concentration of 1% (v/v). 5. Incubate the sample with gentle shaking for 5 min. 6. Harvest the cells by centrifugation at 8000 g for 10 min and resuspend in 4.0 mL of PBS. 7. Add 10% (w/v) dodecyl maltoside to the suspension at a final concentration of 1% (w/v). 8. Incubate the suspension with gentle shaking for 5 min at RT. 9. Remove cell debris by centrifugation at 8000 g for 10 min at RT. 10. Add ammonium sulfate to the suspension at a final concentration of 20% (w/v). 11. Collect the insoluble fraction by centrifugation at 15,000 g for 15 min. 12. Resuspend the pellet in PBS containing 1% (w/v) dodecyl maltoside.

3.4 Observation of Isolated SprB Filaments by TEM

1. Apply the sample from Subheading 3.3, step 12, to an EM grid and incubate the grid for 1 min at RT. 2. Remove liquids from the grid as described in Subheading 3.2, step 3. The blotting procedures are performed in the same way unless otherwise indicated. 3. Apply PBS to the grid. 4. Remove PBS using a filter paper. 5. Apply AM staining solution to the grid and stain the SprB filaments for 1 min. 6. Remove the staining solution completely with a filter paper blotting from the edge of the grid and air dry. 7. Observe the grid as described in Subheading 3.2, step 9. 8. Figure 1b shows a typical example image of the negative stained SprB filaments (see Note 15).

3.5 Preparation of Osmotically Shocked F. johnsoniae Cells

1. Harvest cells from a 1.5 mL volume of a MM cell culture by centrifugation at 5000 g for 1 min at 4 °C. 2. Suspend the cell pellet in 100 μL of ice-cold sucrose solution and place the suspension on ice for 15 min.

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3. Add 1.4 mL of ice-cold ultrapure water and immediately mix gently by inverting the tube for 3 min. 4. Remove intact cells by low-speed centrifugation at 9000 g for 3 min at 4 °C. 5. Collect the osmotically shocked cells from the supernatant by high-speed centrifugation at 20,000 g for 10 min at 4 °C. 6. Suspend the pellet in 50 μL of ice-cold ultrapure water. 3.6 Preparation of Osmotically Shocked S. grandis Cells

1. Add paraformaldehyde solution to 1.5 mL of culture medium at a final concentration of 1.6%, and fix the cells for 30 min at RT (see Note 16). 2. Harvest fixed cells by centrifugation at 5000 g for 1 min at 4 ° C. 3. Suspend the cell pellet in 1.5 mL of fresh marine broth liquid medium. 4. Repeat steps 2 and 3 to remove the fixing solution completely. 5. Harvest fixed cells by centrifugation at 5000 g for 1 min at 4 ° C. 6. Suspend the cell pellet in 1.5 mL of ice-cold MgCl2 solution and mix gently by inverting the tube for 3 min (see Note 17). 7. Remove intact cells by low-speed centrifugation at 2000 g for 3 min at 4 °C. 8. Harvest osmotically shocked cells by high-speed centrifugation at 20,000 g for 5 min at 4 °C. 9. Suspend the pellet in 50 μL of ice-cold MgCl2 solution.

3.7 Observation of the Gliding Machinery of Osmotically Shocked Cells by TEM

1. Apply osmotically shocked cells onto an EM grid and incubate the grid for 1 min at RT. 2. Remove the liquid from the grid as described in Subheading 3.2, step 3. The blotting procedures are performed in the same way unless otherwise indicated. 3. Apply PTA or UA staining solution to the grid and stain the osmotically shocked cells for 1 min. PTA and UA are used for S. grandis and F. johnsoniae, respectively. 4. Remove the staining solution completely with filter paper blotting from the edge of the grid and air dry. 5. Observe the grid as described in Subheading 3.2, step 9 (see Note 18). 6. Figure 2a shows an example image of osmotically shocked F. johnsoniae (see Note 19). 7. Figure 2b shows an example image of the osmotically shocked S. grandis (see Note 20).

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Fig. 2 Electron micrographs of the bundle filament structure disassociating from the osmotically shocked cell. The osmotic pressure causes the cell to burst, and a part of the gliding track associated with outer membrane is peeled off from the cell. (a) Negatively stained osmotically shocked F. johnsoniae. The gliding track and outer membrane are indicated by the black and white arrowhead, respectively. (b) Negatively stained osmotically shocked S. grandis cell. Bars, 200 nm

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Notes 1. The F. johnsoniae type strain ATCC17061 is a model organism for studies on Bacteroidota gliding motility [15]. 2. This is a modification of the original MM containing 3.3 mM Tris–HCl pH 7.5 [16]. 3. S. grandis str. Lewin is a whole-genome sequenced strain [17]. 4. The carbon-coated EM grids have a hydrophobic surface. Glow discharge treatment of the grids is required before use to make the grid surface hydrophilic. 5. Dilute the concentration of the staining solution if the TEM image background is too strong. 6. The PTA staining solution with a neutral pH is suitable for staining structures that are sensitive to acidic conditions. The UA staining solution is acidic (pH 4.0). 7. Typical accelerating voltages for a biological TEM range from 70 to 125 kV. The settings may vary depending on the microscope used. 8. F. johnsoniae cells on a CYE plate remain viable for one week at RT. 9. The middle exponential growth phase of a cell culture should be used for the osmotically shocked cell procedure. The cells in the stationary phase are more resistant to osmotic changes.

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10. S. grandis cells on marine broth plates remained viable for one week at RT. 11. S. grandis has a filamentous cell shape that is approximately 1 μm wide and 5–500 μm long. To prevent cell damage, cells should be cultured with slow shaking. 12. Frozen cells of both species remain viable for at least several months at -80 °C. 13. Keep the grid surface wet. Drying the grid surface during staining may cause structural artifacts and salt crystallization. 14. The adhesin SprB filaments expand from the cell surface (Fig. 1a) [9]. 15. SprB filaments approximately 150 nm long can be observed (Fig. 1b) [9]. 16. The structure of the gliding machinery in S. grandis (Fig. 2b) is fragile and disassembled in water without the glutaraldehyde fixation [2]. 17. At least 40 mM of magnesium is required to stabilize the gliding machinery of S. grandis [10]. Cells burst due to osmotic pressure, but the gliding machinery remains associated with the cell membrane in the presence of magnesium. The 50 mM magnesium ion concentration is similar to that of seawater. 18. The osmotically shocked cells appeared transparent (Fig. 2a, b). Gaps are observed between the outer and inner membranes (Fig. 2a, white arrowhead). In contrast, gaps are not observed in intact cells (Fig. 1a). 19. Bundle filaments associated with the membranes can be observed (Fig. 2a). This structure is absent in gliding motility-deficient mutants in F. johnsoniae (unpublished), suggesting that the structure may be a part of the helical track of the gliding machinery. 20. Bundle filaments covering the cell surface of S. grandis can be observed (Fig. 2b).

Acknowledgments We thank Dr. M. M. Matthews (Okinawa Institute of Science and Technology Graduate University) for English language editing. This work was supported by KAKENHI (17K17085, 19K10083 to SS and 16H06230, 20H05543, 21K07020 to DN) and funds from the Noguchi Institute for DN.

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References 1. Kubori T, Matsushima Y, Nakamura D et al (1998) Supramolecular structure of the Salmonella typhimurium type III protein secretion system. Science 280:602–605. https://doi. org/10.1126/science.280.5363.602 2. Aizawa S-I (2014) The Flagellar World. Academic Press. https://doi.org/10.1016/ C2013-0-06810-7 3. Kubori T, Koike M, Bui XT et al (2014) Native structure of a type IV secretion system core complex essential for Legionella pathogenesis. Proc Natl Acad Sci U S A 111:11804–11809. https://doi.org/10.1073/pnas.1404506111 4. McBride MJ (2001) Bacterial gliding motility: multiple mechanisms for cell movement over surfaces. Ann Rev Microbiol 55:49–75. https://doi.org/10.1146/annurev.micro.55. 1.49 5. Miyata M, Robinson RC, Uyeda TQP et al (2020) Tree of motility – A proposed history of motility systems in the tree of life. Genes Cells 25:6–21. https://doi.org/10.1111/gtc. 12737 6. McBride MJ (2019) Bacteroidetes gliding motility and the type IX secretion system. Microbiol Spectr 7. https://doi.org/10. 1128/microbiolspec.PSIB-0002-2018 7. Hennell JR, Deme JC, Kjær A et al (2021) Structure and mechanism of the proton-driven motor that powers type 9 secretion and gliding motility. Nat Microbiol 6:221–233. https:// doi.org/10.1038/s41564-020-00823-6 8. Liu J, McBride MJ, Subramaniam S (2007) Cell surface filaments of the gliding bacterium Flavobacterium johnsoniae revealed by cryoelectron tomography. J Bacteriol 189:7503– 7 5 0 6 . h t t p s : // d o i . o r g / 1 0 . 1 1 2 8 / J B . 00957-07 9. Nakane D, Sato K, Wada H et al (2013) Helical flow of surface protein required for bacterial gliding motility. Proc Natl Acad Sci U S A

110:11145–11150. https://doi.org/10. 1073/pnas.1219753110 10. Aizawa S (2005) Bacterial gliding motility: visualizing invisible machinery. ASM News 71:71–76 11. Katayama E, Tahara YO, Bertin C, Shibata S (2019) Application of spherical substrate to observe bacterial motility machineries by Quick-Freeze-Replica Electron Microscopy. Sci Rep 9:14765. https://doi.org/10.1038/ s41598-019-51283-w 12. Braun TF, McBride MJ (2005) Flavobacterium johnsoniae GldJ is a lipoprotein that is required for gliding motility. J Bacteriol 187:2628– 2637. https://doi.org/10.1128/JB.187.8. 2628-2637.2005 13. Sato K, Naito M, Yukitake H et al (2010) A protein secretion system linked to bacteroidete gliding motility and pathogenesis. Proc Natl Acad Sci U S A 107:276–281. https://doi. org/10.1073/pnas.0912010107 14. Abby SS, Rocha EPC (2012) The non-flagellar type III secretion system evolved from the bacterial flagellum and diversified into host-cell adapted systems. PLoS Genet 8. https://doi. org/10.1371/journal.pgen.1002983 15. McBride MJ, Kempf MJ (1996) Development of techniques for the genetic manipulation of the gliding bacterium Cytophaga johnsonae. J Bacteriol 178:583–590. https://doi.org/10. 1128/jb.178.3.583-590.1996 16. Nelson SS, Glocka PP, Agarwal S et al (2007) Flavobacterium johnsoniae SprA is a cell surface protein involved in gliding motility. J Bacteriol 189:7145–7150. https://doi.org/10.1128/ JB.00892-07 17. Saw JHW, Yuryev A, Kanbe M et al (2012) Complete genome sequencing and analysis of Saprospira grandis str. Lewin, a predatory marine bacterium. Stand Genom Sci 6:84–93. https://doi.org/10.4056/sigs.2445005

Chapter 22 Live Cell Imaging of Gliding Motility of Flavobacterium johnsoniae Under High-Resolution Microscopy Daisuke Nakane and Satoshi Shibata Abstract Many phylum Bacteroidetes bacteria are motile without either flagella or pili. These cells move on surfaces such as glass or agar, and a motor generates a propulsion force for the cells via a proton motive force across the cytoplasmic membrane. The gliding motility depends on the helical track of cell adhesin along the longer axis of the cell body. Here, we describe live-cell imaging of gliding motility under optical microscopy, as well as an immunofluorescent labeling method for visualizing helical trajectories. Key words Surface-associated motility, Helical track, Adhesin, Flavobacterium, Optical microscopy

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Introduction Many members of the phylum Bacteroidetes family can move smoothly over surfaces [1–3]. These cells move back and forth along their long axes of the cell body, pivoting, flipping, and occasionally rotating at a cell pole (Fig. 1a). This type of bacterial cell movement, known as gliding motility, has evolved as a distinct molecular mechanism for achieving motion without the use of either flagella or pili [4, 5]. In Flavobacterium johnsoniae, adhesins such as SprB are anchored to the outer membrane and repeatedly move from one pole to another along the longer axis of the cell in helical loops to transmit cell propulsion forces (Fig. 2a) [6–8]. The helical track is also used for a type of gliding motility in myxobacteria, known as adventurous or A-motility [9–11], though these two distinct gliding motility systems in F. johnsoniae and myxobacteria may have evolved independently [12, 13]. Although the protein component of the helical track in the cell is still unknown in F. johnsoniae, the lipoprotein GldJ exhibits a helical arrangement within the cell envelope [14]. Cryo-electron tomography and quick-freeze-replica EM visualize patches at the inner surface of

Tohru Minamino et al. (eds.), Bacterial and Archaeal Motility, Methods in Molecular Biology, vol. 2646, https://doi.org/10.1007/978-1-0716-3060-0_22, © The Author(s), under exclusive license to Springer Science+Business Media, LLC, part of Springer Nature 2023

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Fig. 1 Gliding motility of F. johnsoniae cells. (a) Characteristic cell behaviors on a glass surface: back and forth, pivoting/flipping, and rotating at a pole. (b) Cell image under phase-contrast microscopy (left). Cell trajectory for 45 s (right). Sequential images over time are integrated into one image. (c) Sample chamber. Two pieces of double-sided tape were used to assemble two coverslips. The cell suspension was poured into the tunnel. The cells were observed near the double-sided tape as indicated by dashed circles

Fig. 2 Helical trajectories of surface adhesin for gliding motility in F. johnsoniae. (a) Schematic model of corkscrew motion. Surface proteins such as SprB move along helical tracks on the cell surface (upper). Machinery drives the gliding motility (lower). IM: inner membrane. PG: peptidoglycan. OM: outer membrane. Helical track is presented as a short fragment located at the inner side of the outer membrane. (b) Dynamics of SprB in living cells. Kymograph of SprB signaling by immunofluorescent labeling. The x-axis represents the position of the SprB signals, and the y-axis is time. (c) Localization of SprB in a cell

the outer membrane and the intracellular stripe structure, respectively, and these structures may be part of the helical track architecture for SprB movements [15, 16, 34]. The gliding motility of Bacteroidetes is associated with a type IX secretion system (T9SS) [3, 17–19]. The protein complex consisting of GldK, GldL, GldM, and GldN transports large proteins, such as cell-surface adhesins, across the outer membrane [20] and propels them, thereby achieving cell movement powered by proton

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motive force (PMF) across the cytoplasmic membrane [7, 36]. GldK and GldN form a 50 nm diameter ring on the inner side of the outer membrane [21], whereas GldL and GldM span the cytoplasmic membrane [35]. A recent cryo-EM study has revealed that the transmembrane core of GldL and GldM for Flavobacteria gliding motility has a structural organization similar to that of the bacterial flagellar stator complex consisting of MotA and MotB, which acts as a proton channel that couples the proton flow through the channel to torque generation [22–24, 37]. This raises the open question about the molecular mechanism and evolution of the structure of proton-driven motors, as the rotational motion in both biological systems may be generated by the same mechanism [24]. GldL is localized near the axis of rotation of a tethered cell [25], which is the only experimental evidence for motor rotation. More research is expected on how the gliding motor transmits force to the periplasmic side, allowing adhesins to move linearly from pole to pole like the caterpillar track (Fig. 2a). Here, we describe methods to observe gliding motility of F. johnsoniae on glass surfaces (Fig. 1b) and to visualize helical trajectories of surface proteins during cell movement with either immunofluorescent labeling or fluorescent beads in living cells (Fig. 2b).

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Materials Prepare all solutions using ultrapure water and analytical grade reagents. Sterilize the solutions (autoclave for 20 min at 121 °C) prior to use and store all reagents at 4 °C unless indicated otherwise.

2.1 Strain Preparation

1. F. johnsoniae ATCC17061 (see Note 1). 2. CYE liquid medium [26]: 10% [w/v] casitone, 5% yeast extract, and 8 mM MgSO4 in 10 mM Tris–HCl [pH 7.5]. 3. CYE plate: 1% casitone, 0.5% yeast extract, 8 mM MgSO4, and 1.5% agar in 10 mM Tris–HCl [pH 7.5]. 4. Disposable test tube: 15 ml. 5. Incubator set to 25 °C. 6. Shaker. 7. Spectrophotometer. 8. Motility buffer (MB): 20 mM Tris–HCl, pH 7.5, 5 mM MgSO4. 9. Filter paper. 10. Nail polish.

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2.2 Optical Microscopy

1. Inverted microscope. 2. Objective lens for phase-contrast microscopy (see Note 2). 3. Condenser for phase-contrast microscopy. 4. Camera (see Note 3). 5. Optical filter set for fluorescent dye. 6. Epi-illuminator for fluorescence microscopy.

2.3 Glass Chamber Assembly

1. Coverslip: 22 × 40 mm. 2. Coverslip: 18 × 18 mm. 3. Double-sided tape (see Note 4).

2.4 Immunofluorescent Labeling of SprB

1. Primary antibody: Antisera against SprB [6].

2.5 Inhibitor and Probe for Cell Surface Movement

1. CCCP: 50 μM carbonyl cyanide 3-chlorophenylhydrazone in CYE.

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2. Secondary antibody: Fluorescently labeled anti-rabbit IgG (see Note 5).

2. Fluorescent beads: Carboxylate-modified beads with a diameter of 200 nm (see Note 6).

Methods Carry out all procedures at room temperature (RT) unless otherwise specified.

3.1 Culture Conditions

1. Streak the frozen stock of F. johnsoniae cells on a CYE plate (see Note 7). 2. Incubate the plate at 25 °C for at least 24 h (see Note 8). 3. Prepare a test tube with 2 ml CYE liquid medium. 4. Inoculate a single colony of the F. johnsoniae cells from the plate into the test tube. 5. Incubate the test tube at 25 °C with shaking at 180 rpm. 6. Grow the cells to an optical density of 0.5–1.0 at 600 nm, as measured by a spectrophotometer (see Note 9).

3.2 Gliding Motility Observed Under Phase-Contrast Microscopy

1. Assemble a sample chamber with double-sided tape, separating the coverslips with two pieces of double-sided tape to act as a thin channel for observing the sample (Fig. 1c). 2. Transfer the cell culture into the chamber and incubate at RT for 5 min. 3. Add MB to replace the solution in the chamber (see Note 10). 4. Seal both ends of the chamber with nail polish to ensure that the sample does not dry out.

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5. Place the chamber on the microscope stage. 6. Focus on a cell located near the double-sided tape (Fig. 1c) (see Note 11). 7. Capture cell images by a camera and record sequential images at an exposure time of at least 0.1 s. 8. Observe the cells for 20 min (see Notes 12 and 13). 3.3 Visualization of SprB Dynamics with Immunofluorescent Labeling

1. Prepare cells in the chamber, as described in Subheading 3.2, steps 1–3. 2. Dilute the primary antibody into fresh CYE liquid medium at a dilution of 1:100. 3. Add the primary antibody to replace the solution in the chamber (see Notes 10 and 14). 4. Incubate the sample chamber for 5 min at RT. 5. Add fresh CYE medium to replace the solution in the chamber (see Note 10). This step should be repeated at least three times to ensure that the antisera is completely removed. 6. Dilute the secondary antibody into fresh CYE liquid medium at a dilution of 1:100. 7. Add the secondary antibody to replace the solution in the chamber (see Note 10). 8. Incubate the sample chamber for 5 min at RT. 9. Add fresh CYE medium to replace the solution in the chamber (see Note 10). This step should be repeated at least three times to ensure that the secondary antibody is completely removed. 10. Seal both ends of the chamber with nail polish to ensure that the sample does not dry out. 11. Place the chamber on the microscope stage, as described in Subheading 3.2, steps 5–7. 12. Apply the excitation light for the fluorescent dye, and record sequential images at an exposure time of at least 0.1 s. 13. Perform the observations described in Subheading 3.3, steps 11 and 12 within 10 min (see Notes 15 and 16). 14. Open the sequential image file of the SprB-labeled cell and create a kymograph (Fig. 2b) (see Note 17).

3.4 Inhibition of Gliding Motility by CCCP

1. Prepare cells in the chamber, as described in Subheading 3.2, steps 1–3. 2. Place the chamber on the microscope stage. 3. Add CCCP to replace the solution in the chamber (see Notes 10 and 18). 4. Perform the observations described in Subheading 3.2, steps 5–7 within 10 min (see Note 19).

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3.5 Visualization of Helical Movement on the Cell Surface Through Fluorescent Beads

1. Prepare cells in the chamber, as described in Subheading 3.2, steps 1–3. 2. Dilute fluorescent beads into fresh CYE liquid medium at a dilution of 1:100. 3. Add the fluorescent beads to replace the solution in the chamber (see Note 10). 4. Seal both ends of the chamber with nail polish to ensure that the sample does not dry out. 5. Place the chamber on the microscope stage, as described in Subheading 3.2, steps 5–7. 6. Apply the excitation light for the fluorescent dye, and record sequential images at an exposure time of at least 0.1 s.

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Notes 1. F. johnsoniae ATCC17061 (UW101) is the genome sequenced strain, and many genes have been identified to be involved in the gliding motility [27]. F. johnsoniae was formerly called Cytophaga johnsoniae before 1996 [26]. 2. Objective lens is important to see cell behavior under optical microscopy. Example is LUCPLFLN20×PH and UPLFLN 100×O2PH (Olympus) for the images in Figs. 1 and 2, respectively. 3. Use highly sensitive camera to obtain clear fluorescent images under fluorescence microscopy. Example is iXon3 897 or Zyla 4.2 (Andor). 4. Double-sided tape made of nonwoven fabrics such as NWBB-5 (Nichiban) was used. This allowed the aerobic bacteria to be supplied with oxygen in the sample chamber. 5. The primary antibody was prepared from rabbits using recombinant SprB protein [6]. An anti-rabbit IgG secondary antibody conjugated with Cy3 is commercially available as AP132C from Sigma–Aldrich [7]. Use proper optical filter to obtain high signal-to-noise ratio under fluorescence microscopy. Example is an optical filter set Cy3-4040C (Semrock). 6. Carboxylate-modified beads (F8811, Thermo Fisher) were used for cell surface attachment. Other surfaces, such as amine-modified beads, are ineffective for visualizing helical trajectories. 7. The frozen stock was prepared by cell culture at an OD600 of 1.0, mixed with glycerol to a final concentration of 15% [v/v], and stored at -80 °C. 8. The cell plate should be stored at RT and used within a week.

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9. The cells should be prepared from a culture of late exponential phase. Gliding motility is not clear during the stationary phase. The doubling time of F. johnsoniae is approximately 1–2 h under optimal growth conditions. 10. Place a piece of filter paper on one side of the tunnel in the chamber, and add the reagent of interest from the other side. 11. The F. johnsoniae cells exhibit clear gliding motility near the double-sided tape [7]. This is most likely due to the higher concentration of dissolved oxygen. A similar method exists for other species [2, 28]. This procedure should be modified for optimal growth conditions when aerophilic bacteria or anaerobic bacteria are used for observation. 12. A projected image to a camera allows for a clear view of cell behavior (Fig. 1b) [29]. The sequential images of gliding motility should be opened in a software such as ImageJ (https://imagej.nih.gov/ij/), and Z-projection should be applied on the sequential images. Coloring individual frames from red (time zero) to yellow, green, cyan, and finally blue and combining them into one integrated image results in rainbow traces, which helps us to observe multiple cell behaviors at the same time [30]. 13. The cells showed a variety of movements on the glass surface, such as back and forth movement and pivoting/flipping (Fig. 1a). Rotating behavior at one pole is rarely observed, although this was reported 40 years ago [31, 32]. 14. The antisera concentration is a tradeoff between labeling efficiency and motility inhibition [7]. Antisera at concentrations greater than 1/100 dilution reduced the time required to label SprB with antisera. However, higher concentrations of antisera caused the SprB and antisera to aggregate, which inhibited gliding motility and eventually caused the cells to detach from the glass surface. 15. All steps in Subheading 3.3 should be performed quickly for live imaging. Immunofluorescence microscopy of SprB in a chemically fixed cell revealed a large number of dot-like signals in one region of the cell (Fig. 2c). However, these signals gradually combine to form a strong signal in a living cell [7]. 16. Fluorescently labeled SprB shows repeated helical trajectories from one cell pole to the other [29]. However, the chirality of the helical track is uncertain: the trajectory is a left-handed helix under total internal reflection fluorescence microscopy (TIRFM) [7], while it is a right-handed helix under threedimensional tracking of gold nanoparticles [33]. 17. The kymograph depicts the spatial position of SprB over time [7]. The sequential images should be opened in software such as ImageJ. Then, a cell moving parallel to the x-axis should be

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located, the region of cell movement should be cropped, and the height of the region should be resized to one pixel. The resized sequential image should be vertically aligned, and the data should be displayed so that the y-axis represents time (Fig. 2b). 18. CCCP was dissolved in dimethyl sulfoxide (DMSO) at 10 mM and diluted to 10 μM in CYE. CCCP can be used as an uncoupler to inhibit proton motive force. 19. The gliding motility of the cells stops within a few seconds. The inhibitory effect of SprB movement can be observed by using the sample chamber described in Subheading 3.3.

Acknowledgments This work was supported by KAKENHI (16H06230, 20H05543, 21K07020, 22H05066 to DN, and 17K17085, 19K10083 to SS) and funds from the Noguchi Institute to DN. We thank Mark McBride for kindly gifting the antiserum against SprB. References 1. McBride MJ (2001) Bacterial gliding motility: multiple mechanisms for cell movement over surfaces. Annu Rev Microbiol 55:49–75. https://doi.org/10.1146/annurev.micro.55. 1.49 2. McBride MJ, Zhu Y (2013) Gliding motility and Por secretion system genes are widespread among members of the phylum Bacteroidetes. J Bacteriol 195:270–278. https://doi.org/10. 1128/jb.01962-12 3. McBride MJ, Sandkvist M, Cascales E et al (2019) Bacteroidetes gliding motility and the type IX secretion system. Microbiol Spectr 7: 7.1.15. https://doi.org/10.1128/micro biolspec.PSIB-0002-2018 4. Miyata M, Robinson RC, Uyeda TQP et al (2020) Tree of motility – a proposed history of motility systems in the tree of life. Genes Cells 25:6–21. https://doi.org/10.1111/gtc. 12737 5. Wadhwa N, Berg HC (2022) Bacterial motility: machinery and mechanisms. Nat Rev Microbiol. https://doi.org/10.1038/ s41579-021-00626-4 6. Nelson SS, Bollampalli S, McBride MJ (2008) SprB is a cell surface component of the Flavobacterium johnsoniae gliding motility machinery. J Bacteriol 190:2851–2857. https://doi. org/10.1128/jb.01904-07

7. Nakane D, Sato K, Wada H et al (2013) Helical flow of surface protein required for bacterial gliding motility. Proc Natl Acad Sci U S A 110:11145–11150. https://doi.org/10. 1073/pnas.1219753110 8. Wada H, Nakane D, Chen H-Y (2013) Bidirectional bacterial gliding motility powered by the collective transport of cell surface proteins. Phys Rev Lett 111:248102. https://doi.org/ 10.1103/PhysRevLett.111.248102 9. Mignot T, Shaevitz JW, Hartzell PL et al (2007) Evidence that focal adhesion complexes power bacterial gliding motility. Science 315: 853–856. https://doi.org/10.1126/science. 1137223 10. Nan B, Chen J, Neu JC et al (2011) Myxobacteria gliding motility requires cytoskeleton rotation powered by proton motive force. Proc Natl Acad Sci U S A 108:2498–2503. https://doi. org/10.1073/pnas.1018556108 11. Faure LM, Fiche J-B, Espinosa L et al (2016) The mechanism of force transmission at bacterial focal adhesion complexes. Nature 539: 5 3 0 – 5 3 5 . h t t p s : // d o i . o r g / 1 0 . 1 0 3 8 / nature20121 12. Nan B, McBride Mark J, Chen J et al (2014) Bacteria that glide with helical tracks. Curr Biol 24:R169–R173. https://doi.org/10.1016/j. cub.2013.12.034

Live Imaging of Gliding Motility of Flavobacterium 13. Nan B, Zusman DR (2016) Novel mechanisms power bacterial gliding motility. Mol Microbiol 101:186–193. https://doi.org/10.1111/ mmi.13389 14. Braun TF, McBride MJ (2005) Flavobacterium johnsoniae GldJ is a lipoprotein that is required for gliding motility. J Bacteriol 187:2628–2637. https://doi.org/10.1128/jb.187.8.2628-2637. 2005 15. Liu J, McBride MJ, Subramaniam S (2007) Cell surface filaments of the gliding bacterium Flavobacterium johnsoniae revealed by cryoelectron tomography. J Bacteriol 189:7503– 7506. https://doi.org/10.1128/jb.00957-07 16. Katayama E, Tahara YO, Bertin C et al (2019) Application of spherical substrate to observe bacterial motility machineries by QuickFreeze-Replica Electron Microscopy. Sci Rep 9:14765. https://doi.org/10.1038/s41598019-51283-w 17. Sato K, Naito M, Yukitake H et al (2010) A protein secretion system linked to bacteroidete gliding motility and pathogenesis. Proc Natl Acad Sci U S A 107:276–281. https://doi. org/10.1073/pnas.0912010107 18. McBride MJ, Nakane D (2015) Flavobacterium gliding motility and the type IX secretion system. Curr Opin Microbiol 28:72–77. https://doi.org/10.1016/j.mib.2015.07.016 19. Trivedi A, Gosai J, Nakane D et al (2022) Design principles of the rotary type 9 secretion system. Front Microbiol 13:845563. https:// doi.org/10.3389/fmicb.2022.845563 20. Shrivastava A, Johnston JJ, van Baaren JM et al (2013) Flavobacterium johnsoniae GldK, GldL, GldM, and SprA are required for secretion of the cell surface gliding motility adhesins SprB and RemA. J Bacteriol 195:3201–3212. https://doi.org/10.1128/jb.00333-13 21. Gorasia DG, Veith PD, Hanssen EG et al (2016) Structural insights into the PorK and PorN components of the Porphyromonas gingivalis type IX secretion system. PLoS Pathog 12:e1005820. https://doi.org/10.1371/jour nal.ppat.1005820 22. Deme JC, Johnson S, Vickery O et al (2020) Structures of the stator complex that drives rotation of the bacterial flagellum. Nat Microbiol 5:1553–1564. https://doi.org/10.1038/ s41564-020-0788-8 23. Santiveri M, Roa-Eguiara A, Ku¨hne C et al (2020) Structure and function of stator units of the bacterial flagellar motor. Cell 183:244– 257.e216. https://doi.org/10.1016/j.cell. 2020.08.016 24. Hennell James R, Deme JC, Kjær A et al (2021) Structure and mechanism of the

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and gliding motility in Flavobacterium. PLOS Biol 20:e3001443. https://doi.org/10.1371/ journal.pbio.3001443 37. Hennell James R, Deme JC, Hunter A et al (2022) mBio 13:e0026722. https://doi.org/ 10.1128/mbio.00267-22

Chapter 23 Social Motility Assays of Flavobacterium johnsoniae Chikara Sato and Keiko Sato Abstract Flavobacterium johnsoniae cells move rapidly over solid surfaces by gliding motility. The collective migration of F. johnsoniae on the surfaces results in the formation of spreading colonies. Colony spreading is influenced by adhesin components on the cell surface and the concentrations of agar and glucose. For example, on nutrient-poor agar media, film-like, round spreading colonies are formed. F. johnsoniae displays at least two types of migration: small cell cluster movements leading to concentric colonies and individual cell movements leading to dendritic colonies. The methods for observing colony morphology are described in this chapter. Key words Social motility, Gliding motility, Colony spreading, Surface adhesin

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Introduction Flavobacterium johnsoniae is an aerobic, gram-negative, rod-shaped bacterium that moves rapidly over solid surfaces by gliding motility and forms film-like round spreading colonies on nutrient-poor agar media [1, 2]. Gliding motility is common in several other species of the Bacteroidetes phylum [3]. Flavobacterium columnare and Flavobacterium psychrophilum are widely distributed fish pathogens and move over solid surfaces by gliding motility, forming biofilms when colonizing the gills of the fish [4–6]. Genetic analyses of F. johnsoniae have revealed that the gld (gliding) genes (gldA, gldB, gldD, gldF, gldG, gldH, gldI, gladJ, gldK, gldL, gldM, gldN, and gldO) are required for gliding and that the spr (spreading) genes (sprA, sprB, sprC, sprD, sprE, sprF, and sprT) are involved in colony spreading [7–14]. Therefore, these gld and spr mutants influence gliding and form non-spreading colonies on agar medium. Electron microscopy with negative staining has shown that the cells of spreading wild-type (WT) colonies are embedded in a selfsecreted matrix containing vesicles and a filamentous network, which is the indicative of biofilm formation [15]. The cell density

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at the bottom of the colony is higher than that in the middle. At the leading edge of the colony, small cell clusters are followed by many cells connected by filaments [15]. Near the leading edge of the colony, the cells make head-to-tail and/or side-to-side contacts and are surrounded by a poorly stained filamentous matrix. However, in non-spreading sprB colonies, the cells are tightly packed [15–17]. The colony spreading of F. johnsoniae is affected by the agar and glucose concentrations [16]. SprB is not required for spreading dendritic colonies on soft agar-containing glucose (adhesinindependent colony spreading). Both WT and the sprB mutant form dendritic colonies on soft agar-containing glucose. The cells move in all directions and spread toward the outer edge of the colony [16]. The colonies spread during the initial growth-dependent phase, which is followed by the gliding motility-dependent phase. Thus, adhesin-dependent and adhesinindependent colony spreading are both influenced by the gliding machinery [16]. Microstructural information was obtained by atmospheric scanning electron microscopy (ASEM) and transmission EM, respectively. ASEM is an inverted SEM that enables observation of a wet sample such as cultured cells and biofilms from below while an optical microscopy simultaneously observes it from above [18–22].

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Materials Prepare all solutions using ultrapure water and analytical grade reagents and then autoclave them at 121  C for 20 min.

2.1 Bacterial Strains and Plasmids

1. Target strain: F. johnsoniae wild-type Cj1827 [23]. 2. Competent cells of E. coli S17-1 λpir [24]. 3. pRR51: a mobilizable suicide vector [23]. 4. pNS1: Bacteroides–E. coli shuttle vector [25]. 5. pFj29: a plasmid encoding a green fluorescent protein, GFPmut3, under a control of the Flavobacterium ompA promoter [26].

2.2 Bacterial Culture Media

1. LB medium: Dissolve 1.0 g of tryptone, 0.5 g of yeast extract, 0.5 g of NaCl and 1.5 g of Bacto Agar in 100 mL of water and sterilize by autoclaving. For the selection of ampicillin-resistant Escherichia coli strains, add ampicillin to the molten agar at a concentration of 100 μg/mL. 2. Casitone-yeast extract (CYE) medium: Dissolve 1.0 g of Casitone, 0.5 g of yeast extract, and 1.5 g of Bacto Agar in 100 mL of water and sterilize by autoclaving.

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3. CYE agar plate: Dissolve 1.0 g of Casitone, 0.5 g of yeast extract, and 1.5 g of Bacto Agar in 100 mL of water and sterilize by autoclaving. For the selection and maintenance of erythromycin- and streptomycin-resistant F. johnsoniae strain, add erythromycin (Em) and streptomycin (Sm) to the molten agar at a final concentration of 100 μg/mL each. 4. CYE-Ca agar plate: For conjugation of E. coli and F. johnsoniae, spread 100 μL of 1 M CaCl2 on the CYE agar plate 30 min before use. 2.3 Bacterial Media for Colony Morphology Observation

1. PY2 agar plate: Dissolve 0.2 g of peptone, 0.5 g of yeast extract, and 1.0 g of Bacto Agar in 100 mL of water and sterilize by autoclaving. 2. 1.5 M glucose solution: Dissolve 27.0 g of glucose in 100 mL of water and sterilize by autoclaving. 3. Peptone yeast glucose (PYG) soft agar plate: dissolve 0.2 g of peptone, 0.5 g of yeast extract, and 0.3 g of Ina Agar (Ina Food Industry Co., Ltd, Japan) in 100 mL of water and sterilize by autoclaving. Add 1 mL of 1.5 M glucose solution to the agar at a final concentration of 15 mM (0.3% (w/v) agar-PYG (15 mM)).

2.4 Construction of gld or spr Deletion Mutant

1. Freeze Throw Buffer (FTB): 10 mM PIPES, 15 mM CaCl2, 250 mM KCl, 55 mM MnCl2. Dissolve 0.6 g of PIPES, 0.44 g of CaCl2.2H2O, 3.72 g of KCl in 190 mL of water. Adjust pH between 6.7 and 6.8 using KOH. Add 2.18 g of MnCl2.4H2O and water to a final volume of 200 mL. Sterilize by a filter (0.22 μm). 2. TC buffer: 10 mM Tris–HCl containing 8 mM CaCl2, pH 7.3. 3. PrimeSTAR® Max DNA Polymerase (Takara Bio): A highfidelity PCR kit. 4. Restriction enzymes: BamHI, SalI, NotI and SphI. 5. Optical microscopy (e.g., Olympus BX50F). 6. Andor iQ3. 7. EMCCD camera (e.g., ANDOR iXon3). 8. Micro cover glass.

2.5 Carbon-Grid Stamp-Peel Method [17]

1. F. johnsoniae strains. 2. PY2 agar plate. 3. 10 mM Tris–HCl, pH 7.4. 4. Centrifuge. 5. Colony spread on agar medium plate.

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Fig. 1 ASEM dish. A silicon chip with eight SiN thin-film window is embedded in the bottom center of the ASEM specimen holder dish. The body of the ASEM dish is made of polystyrene and the dish can be used for cell culture in an incubator

6. Stamp-Peel with a glow-discharged flat carbon film on a copper mesh grid. 7. 2.0% (w/v) uranyl acetate. 2.6 Atmospheric Scanning Electron Microscopy (ASEM) [16]

1. Cells cultured on ASEM dishes (see Fig. 1) are fixed with 1% (w/v) paraformaldehyde (PFA) and 3.5% (w/v) glutaraldehyde in 0.1 M phosphate buffer (PB), pH 7.4. 2. Cells are stained with the following NCMIR method (3–9) or positively charged nanogold labeling/PTA staining (10–12). 3. ClairScope ASEM system (JASM-6200, JEOL, Ltd., Tokyo, Japan). 4. ASEM dishes. 5. 1% (w/v) paraformaldehyde (PFA). 6. 3.5% (w/v) glutaraldehyde in 0.1 M phosphate buffer (PB), pH 7.4. 7. 0.15 M cacodylate buffer containing 2 mM calcium chloride (pH 7.4). 8. 0.15 M cacodylate buffer with 1.5% (w/v) potassium ferricyanide. 9. 2% (w/v) aqueous osmium tetroxide (OsO4). 10. 1% (w/v) thiocarbohydrazide (TCH). 11. 2.0% (w/v) uranyl acetate. 12. 0.4% (w/v) lead citrate. 13. Positively charged Nanogold (e.g., Nanoprobes). 14. GoldEnhance-EM. 15. 2% (w/v) Phosphotungstic acid (PTA).

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Fig. 2 ClairScope ASEM system (JASM-6200). The optical microscope is located on the opposite side of the inverted SEM. An ASEM dish is placed at the top end of the inverted SEM

16. ClairScope ASEM system (JASM-6200, JEOL, Ltd., Tokyo, Japan) (see Fig. 2). 2.7 TEM of F. johnsoniae Spreading Colonies (Biofilms) [15–17]

1. 2.5% (w/v) glutaraldehyde in 0.1 M phosphate buffer (PB), pH 7.4. 2. 1% (w/v) Osmium tetroxide (OsO4) in double distilled water (DDW). 3. Gradient series of ethanol (50%, 70%, 80%, 90%, 95%, and 100%) (v/v). 4. QY-1 (n-Butyl glycidyl ether). 5. Gradient series of QY1:Epon812 (1:1, 1:2, 1:3). 6. Epon 812 (e.g., Nisshin-EM Co., Ltd). 7. Microtome (e.g., Leica Ultracut UCT microtome). 8. EM grids. 9. Uranyl acetate. 10. Lead citrate. 11. Transmission electron microscope (e.g., H7600: Hitachi, Tokyo, Japan).

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Methods

3.1 Construction of Targeting Vector Plasmid

1. Amplify the 2.0 kb upstream and 2.0 kb downstream DNA regions of the target gene from the chromosomal DNA of F. johnsoniae UW 101 by PCR using primer pairs, the Fjoh gene number-U-F-BamHI and Fjoh gene number-U-R-SalI primers and the Fjoh gene number-D-F-SalI and Fjoh gene

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number-D-R-SphI primers, respectively, where “U” indicates upstream, “F” indicates forward, “D” indicates downstream, and “R” indicates reverse (see Note 1). 2. Digest the upstream and downstream regions with BamHI and SalI and with SalI and SphI, respectively. Ligate both digested products into the pRR51 vector that has been digested with BamHI and SphI to yield a plasmid named pFM01. 3.2 Introduction of the Targeting Vector Plasmid into E. coli

1. Preparation of competent cells of E. coli S17-1 λpir [24]: culture E. coli S17-1 λpir in LB broth. Collect the E. coli S17-1 λpir cells by centrifugation at 1000  g for 10 min at 4  C. Suspend the cell pellets in FTB and incubate on ice for 10 min. Centrifuge the cells and suspend them in FTB. 2. Mix pFM01 (usually 100 ng) with 100 μL of competent E. coli S17-1 λpir in a tube. 3. Incubate the mixture on ice for 10 min. 4. Incubate the mixture at 42  C for 60 s. 5. Put the tubes back on ice for 2 min. 6. Place all of the culture onto a LB agar plate containing 100 μg/ mL ampicillin. 7. Incubate the plates at 37  C overnight.

3.3 Construction of a F. johnsoniae Deletion Mutant Strain

1. Culture E. coli S17-1 λpir transformed with the plasmid pFM01 in 1 mL of LB medium containing 100 μg/mL ampicillin overnight. 2. Culture F. johnsoniae Cj1827 in 1 mL of CYE broth at 30  C overnight. 3. Wash the cells with fresh LB medium without antibiotics and centrifugation at 1500  g for 10 min, and discard the supernatant. This step is repeated twice. 4. Mix the cultures of 1 mL of plasmid-retained E. coli S17-1 λpir and 1 mL of the F. johnsoniae Cj1827. 5. Centrifuge the mixture at 1000  g for 10 min. 6. Spot the concentrated mixture on the CYE-Ca agar plates and incubate the plates at 30  C for 24 h (see Note 2). 7. Suspend the bacteria in the TC buffer, and seed on the CYE-Em agar plate. 8. Culture for 2 days at 30  C to form colonies. 9. Pick a single colony, inoculate it into 1 mL of CYE medium and culture at 30  C overnight (see Note 3).

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10. Seed on the CYE-Sm agar plate. 11. Culture for 2 days at 30  C to form colonies. 3.4 Construction of the Shuttle Vector to Produce a Complemented Strain of F. johnsoniae

1. Amplify the entire responsible gene region from the chromosomal DNA by PCR.

3.5 Transformation of E. coli S17-1λ with the Shuttle Vector Plasmid

1. Mix pFC01 (usually 100 ng) into 100 μL of competent E. coli S17-1 λpir in a tube.

2. Digest the amplified DNA fragments with BamHI and NotI and insert into the corresponding restriction sites of the pNS1 vector to yield the pFC01 shuttle vector.

2. Incubate the mixture on ice for 10 min. 3. Incubate the mixture at 42  C for 60 s. 4. Put the tubes back on ice for 2 min. 5. Place all of the culture onto an LB agar plate containing 100 μg/mL ampicillin. 6. Incubate the plates at 37  C overnight.

3.6 Conjugative Transfer to Produce a Complemented Strain of F. johnsoniae

1. Culture E. coli S17-1 λpir harboring the pFC01 plasmid in 1 mL of LB medium containing 100 μg/mL ampicillin overnight. 2. Wash the cells with a fresh LB medium without antibiotics, centrifuge at 1500  g for 10 min, and discard the supernatant. This step is repeated twice. 3. Culture F. johnsoniae deficient mutant in 1 mL of the CYE broth at 30  C overnight. 4. Mix 1 mL of E. coli S17-1 λpir harboring pFC01 and 1 mL of the F. johnsoniae mutant. 5. Centrifuge the mixture at 1000  g for 10 min. 6. Spot the concentrated mixture on the CYE-Ca agar plate and culture at 30  C for 24 h (see Note 2). 7. Suspend the bacteria in the TC buffer, and seed on the CYEEm agar plate. 8. Culture for 2 days at 30  C.

3.7 Construction of Green Fluorescent Protein (GFP)-Marked Strains of Flavobacterium sp.

1. Culture E. coli S17-1 λpir carrying the pFj29 plasmid in 1 mL of LB medium containing 100 μg/mL ampicillin overnight. Wash bacterial cells with a fresh LB medium without antibiotics, centrifuge at 1500  g for 10 min and discard the supernatant. This step is repeated twice. 2. Culture Flavobacterium sp. in 1 mL of the CYE broth at 30  C overnight.

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3. Mix 1 mL of E. coli S17-1 λpir harboring the pFj29 plasmid and 1 mL of the Flavobacterium sp. strains. 4. Centrifuge the mixture at 1000  g for 10 min. 5. Spot the concentrated mixture on the CYE-Ca agar plate and culture at 30  C for 24 h. 6. Suspend the bacteria in the TC buffer, and seed on the CYEEm agar plate. 7. Culture for 2 days at 30  C. 3.8 Colony Spreading (See Note 4)

1. Culture F. johnsoniae in the CYE medium at 25  C overnight. 2. Harvest F. johnsoniae cells by centrifugation at 1500  g for 10 min at room temperature and discard the supernatant. 3. Suspend the cells in 10 mM Tris–HCl, pH 7.4. 4. Collect the cells by centrifugation at 1500  g for 10 min, and discard the supernatant. This step is repeated. 5. Resuspend F. johnsoniae cell in 1 ml of 10 mM Tris–HCl, pH 7.4. 6. Add the cells expressing GFP to the bacterial solution at a concentration of 1% (v/v). 7. Spot the bacterial cells on either PY2 or PYG agar plate and incubate the plates at 25  C for 5 days (1 μL/spot). 8. Observe colonies by time-lapse fluorescence microscopy (see Note 5).

3.9 Carbon-Grid Stamp-Peel Method [17]

1. Culture F. johnsoniae in the CYE medium at 25  C overnight. 2. Harvest cells by centrifugation at 1500  g for 10 min at room temperature and discard the supernatant. 3. Wash the cells with 10 mM Tris–HCl, pH 7.4, centrifuge at 1500  g for 10 min and discard the supernatant. This step is repeated twice. 4. Resuspend F. johnsoniae cells in 1 mL of 10 mM Tris–HCl, pH 7.4. 5. Spot the cells on the PY2 agar plate and culture at 25  C for 5 days (1 μL/spot). 6. Lightly press a glow-discharged carbon film/TEM mesh grid onto the colony surface (see Fig. 3). 7. Place sample side of the grid onto a drop of 2.0% (w/v) uranyl acetate for 30 s twice and dry in the air. 8. Observe samples at magnification ranging from 1000 to 50,000 using a transmission electron microscope at an acceleration voltage of 100 kV.

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Fig. 3 Grid Stamp-Peel TEM method. Schematic diagram the stamping of a colony onto a thin-carbon TEM grid. The surface layer of the colony is transferred to the carbon grid by pressing (stamping) the grid 3.10 Atmospheric Scanning Electron Microscopy (ASEM) [16]

1. Cells cultured on ASEM dishes (see Fig. 1) are fixed with 1% (w/v) paraformaldehyde (PFA) and 3.5% (w/v) glutaraldehyde in 0.1 M phosphate buffer (PB), pH 7.4.

3.11 NCMIR Staining Method for ASEM [16]

1. Fix spreading colonies on the agar medium with 1% PFA (w/v) and 3.5% (w/v) glutaraldehyde in 0.1 M PB (pH 7.4) for 30 min at room temperature.

2. Cells are stained with the following NCMIR method 3.11 (2–13) or positively charged nanogold labeling/PTA staining 3.12 (3–6).

2. Wash the fixed spreading colonies with 0.15 M cacodylate buffer containing 2 mM calcium chloride (pH 7.4). 3. Cut out the colony on the agar layer (5 mm  5 mm) from the agar plate. 4. Rinse the spreading colonies with 0.15 M cacodylate buffer containing 2 mM CaCl2. 5. Fix/stain the spreading colonies with 0.15 M cacodylate buffer supplemented with 1.5% (w/v) potassium ferricyanide and 2% (w/v) aqueous osmium tetroxide (OsO4) at room temperature for 20 min. 6. Wash the spreading colonies with DDW. 7. Incubate the spreading colonies with filtered 1% (w/v) thiocarbohydrazide at room temperature for 20 min. 8. Rinse the spreading colonies with DDW. 9. Stain the spreading colonies with 2% (w/v) aqueous OsO4 at room temperature for 30 min. 10. Rinse the spreading colonies with DDW. 11. Stain the spreading colonies with 2% (w/v) uranyl acetate in DDW at 4  C overnight.

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12. Rinse the spreading colonies with DDW. 13. Stain the spreading colonies with 0.4% (w/v) lead citrate at room temperature for 2 min. 14. To image the top of the colony, invert the block of colonies and place it on the SiN film of the ASEM dish. 3.12 Labeling with Charged Nanogold [16]

1. Culture F. johnsoniae on the SiN film window of ASEM dishes. 2. Fix cultured bacterial cells with 1% (w/v) PFA and 3.5% (w/v) glutaraldehyde in 0.1 M PB (pH 7.4) for 30 min at room temperature. 3. Incubate the fixed bacteria on an ASEM dish with a 6 μM solution of positively charged 1.4 nm Nanogold particles for 20 min at room temperature. 4. Wash the bacteria with DDW. 5. To increase the size of the gold particles, carry out gold enhancement using GoldEnhance-EM (Nanoprobes) for 10 min at room temperature. 6. Wash the bacteria with DDW.

3.13 ASEM Imaging [16]

3.14 TEM Imaging of F. johnsoniae Spreading Colonies (Biofilms) [15–17]

Capture ASEM images using the ClairScope ASEM system (JASM6200, JEOL, Ltd., Tokyo, Japan) at an acceleration voltage of 20–30 kV. 1. Fix biofilms with 2.5% (w/v) glutaraldehyde in PB at room temperature for 1 h. 2. Rinse the biofilms with 0.1 M PB (pH 7.4). 3. Stain the samples with 1% (w/v) OsO4 in PB at 4  C for 1 h. 4. Rinse the spreading colonies with 0.1 M PB (pH 7.4). 5. Dehydrate the fixed colonies through a gradient series of ethanol at room temperature. 6. Infiltrate the colonies with QY1 for 15 min. 7. Infiltrate the colonies with Gradient series of QY1:Epon812. 8. Embed the colonies in Epon812. 9. Cut colonies parallel to the colony spreading direction and perpendicular to the agar medium surface into 70 or 400 nm thick ultrathin sections. 10. Mount the ultrathin sections on EM grids. 11. Use the unstained sections directly for TEM, or stain the sections with uranyl acetate and lead citrate. 12. Capture EM images by TEM operating at 80–100 kV.

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Notes 1. Homologous recombination requires the insertion of 2000 bp of upstream and downstream regions of the target gene into the target plasmid. 2. Transfer of plasmids between bacteria via a conjugative transfer system requires cell contact and DNA metabolism between donor and recipient cells. When E. coli and F. johnsoniae are co-cultured on an agar plate, the bacterial pellet forms a spot in one place instead of seeding. 3. Integrant colonies are screened by colony-PCR using a pair of ermF-primers (ermF: erythromycin resistance gene); ermF-F: atgacaaaaaagaaattgc, ermF-R: ctacgaaggatgaaattt. 4. The colony spreading of F. johnsoniae is significantly affected by the agar and glucose concentrations. To observe the surface adhesin SprB-dependent or SprB-independent colony spreading, washed F. johnsoniae cells are inoculated on 1% (w/v) agar PY2 medium or 0.3% (w/v) agar PY2 medium supplemented with 15 mM glucose (PYG), respectively. To visualize the movement of individual bacterial cells at the colony edge, a 1: 100 mixture of WT cells with and without cytoplasmic GFP expression is inoculated on an agar plate. 5. Cells expressing GFP are added to the inoculated bacterial solution at a concentration of 1% (v/v), and the movement of the bacterial cells at the colony edge is observed. Long lines of GFP-producing cells are apparent within the non-spreading colonies.

References 1. Zhu Y, McBride MJ (2014) Deletion of the Cytophaga hutchinsonii type IX secretion system gene sprP results in defects in gliding motility and cellulose utilization. Appl Microbiol Biotechnol 98:763–775 2. Agarwal S, Hunnicutt DW, McBride MJ (1997) Cloning and characterization of the Flavobacterium johnsoniae (Cytophaga johnsonae) gliding motility gene, gldA. Proc Natl Acad Sci U S A 94:12139–12144 3. McBride MJ, Zhu Y (2013) Gliding motility and Por secretion system genes are widespread among members of the phylum bacteroidetes. J Bacteriol 195:270–278 4. Decostere A, Haesebrouck F, Charlier G et al (1999) The association of Flavobacterium columnare strains of high and low virulence with gill tissue of black mollies (Poecilia sphenops). Vet Microbiol 67:287–298

˜ o-Herrera R (2017) Dif5. Levipan HA, Avendan ferent phenotypes of mature biofilm in Flavobacterium psychrophilum share a potential for virulence that differs from planktonic state. Front Cell Infect Microbiol 7:76 6. Kondo M, Kawai K, Kurohara K et al (2002) Adherence of Flavobacterium psychrophilum on the body surface of the ayu Plecoglossus altivelis. Microbes Infect 4:279–283 7. Hunnicutt DW, Kempf MJ, McBride MJ (2002) Mutations in Flavobacterium johnsoniae gldF and gldG disrupt gliding motility and interfere with membrane localization of GldA. J Bacteriol 184:2370–2378 8. Hunnicutt DW, McBride MJ (2000) Cloning and characterization of the Flavobacterium johnsoniae gliding-motility genes gldB and gldC. J Bacteriol 182:911–918

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9. McBride MJ, Braun TF (2004) GldI is a lipoprotein that is required for Flavobacterium johnsoniae gliding motility and chitin utilization. J Bacteriol 186:2295–2302 10. Braun TF, McBride MJ (2005) Flavobacterium johnsoniae GldJ is a lipoprotein that is required for gliding motility. J Bacteriol 187:2628– 2637 11. Braun TF, Khubbar MK, Saffarini DA et al (2005) Flavobacterium johnsoniae gliding motility genes identified by mariner mutagenesis. J Bacteriol 187:6943–6952 12. Rhodes RG, Nelson SS, Pochiraju S et al (2011) Flavobacterium johnsoniae sprB is part of an operon spanning the additional gliding motility genes sprC, sprD, and sprF. J Bacteriol 193:599–610 13. Nakane D, Sato K, Wada H et al (2013) Helical flow of surface protein required for bacterial gliding motility. Proc Natl Acad Sci U S A 110:11145–11150 14. Sato K, Naito M, Yukitake H et al (2010) A protein secretion system linked to bacteroidete gliding motility and pathogenesis. Proc Natl Acad Sci U S A 107:276–281 15. Sato K, Naya M, Hatano Y et al (2021) Biofilm spreading by the adhesin-dependent gliding motility of Flavobacterium johnsoniae. 1. Internal structure of the biofilm. Int J Mol Sci 22: 1894 16. Sato K, Naya M, Hatano Y et al (2021) Colony spreading of the gliding bacterium Flavobacterium johnsoniae in the absence of the motility adhesin SprB. Sci Rep 11:697 17. Sato K, Naya M, Hatano Y et al (2021) Biofilm spreading by the adhesin-dependent gliding motility of Flavobacterium johnsoniae: 2. Role of filamentous extracellular network and cell-to-cell connections at the biofilm surface. Int J Mol Sci 22(13):6911 18. Sakai E, Sato M, Memtily M et al (2021) Liquid-phase ASEM imaging of cellular and structural details in cartilage and bone formed

during endochondral ossification: Keap1deficient osteomalacia. Sci Rep 11:5722 19. Sugimoto S, Okuda K, Miyakawa R et al (2016) Imaging of bacterial multicellular behaviour in biofilms in liquid by atmospheric scanning electron microscopy. Sci Rep 6:25889 20. Okuda KI, Nagahori R, Yamada S et al (2018) The composition and structure of biofilms developed by Propionibacterium acnes isolated from cardiac pacemaker devices. Front Microbiol 9:182 21. Sato C, Yamazaki D, Sato M (2019) Calcium phosphate mineralization in bone tissues directly observed in aqueous liquid by atmospheric SEM (ASEM) without staining: microfluidics crystallization chamber and immuno-EM. Sci Rep 9:7352 22. Naya M, Sato C (2020) Pyrene excimer-based fluorescent labeling of neighboring cysteines by protein dynamics: ASEM-induced thiolene click reaction for high spatial resolution CLEM. Int J Mol Sci 21:7550 23. Rhodes RG, Pucker HG, McBride MJ (2011) Development and use of a gene deletion strategy for Flavobacterium johnsoniae to identify the redundant gliding motility genes remF, remG, remH, and remI. J Bacteriol 193: 2418–2428 24. Simon R, Priefer U, Puhler A (1983) A broad host range mobilization system for in vivo genetic engineering: transposon mutagenesis in gram negative bacteria. Biotechnology 1: 784–791 25. Imamura K, Sato K, Narita Y et al (2018) Identification of a major glucose transporter in Flavobacterium johnsoniae: inhibition of F. johnsoniae Colony spreading by glucose uptake. Microbiol Immunol 62:507–516 26. Chen S, Kaufman MG, Bagdasarian M et al (2010) Development of an efficient expression system for Flavobacterium strains. Gene 458: 1–10

Chapter 24 Visualization of Peptidoglycan Structures of Escherichia coli by Quick-Freeze Deep-Etch Electron Microscopy Yuhei O. Tahara and Makoto Miyata Abstract Peptidoglycan (PG) is an essential component of the bacterial cell wall that protects the cell from turgor pressure and maintains its shape. In diderm (gram-negative) bacteria, such as Escherichia coli, the PG layer is flexible with a thickness of a 2–6 nm, and its visualization is difficult due to the presence of the outer membrane. The quick-freeze deep-etch replica method has been widely used for the visualization of flexible structures in cell interior, such as cell organelles and membrane components. In this technique, a platinum replica on the surface of a specimen fixed by freezing is observed using a transmission electron microscope. In this chapter, we describe the application of this method for visualizing the E. coli PG layer. We expect that these methods will be useful for the visualization of the PG layer in diverse bacterial species. Key words Quick-freeze deep-etch replica, Peptidoglycan, Electron microscopy, Cell wall, Diderm bacteria

1

Introduction Majority of bacteria have a cell wall composed of peptidoglycan (PG). The PG layer maintains cell morphology, prevents osmotic rupture, and provides scaffolds for motile machineries [1–5]. As the PG layer is essential for survival, it is also a crucial target for the host immune system and antibiotics [6, 7]. It is essential also for many bacterial motility systems including flagella and type IV pili, because they use PG as the scaffolds of motility machineries [5]. Interestingly, in adventurous motility (A-motility) of Myxococcus and Bacteroidetes gliding motility, the force generated around the cell membrane is somehow transmitted to the surface of the outer membrane across the PG layer [8, 9]. The PG layer has a weblike structure consisting of a glycan chain of two alternatively linked sugars and a cross-linking peptide chain that provides a robust, flexible, and light shell structure. However, the weblike structure

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makes them unsuitable for observation by transmission electron microscopy (TEM). The quick-freeze deep-etch replica electron microscopy (QFDE-EM) method was used initially to visualize synaptic transmission processes and then used subsequently to visualize other biological phenomena in diverse fields [10, 11]. In this method, the sample is frozen in less than 1 ms by pressing it against a metal block chilled with liquid helium or liquid nitrogen, resulting in much faster fixation than using chemical methods. The frozen sample is exposed by fracturing and etching and is shadowed with platinum. Metal shadowing provides high-contrast and three-dimensional images. This is of significant advantage, particularly for the visualization of low-density and flexible structures. Almost all bacterial species can be classified into two types based on their peripheral structures [12]. Monoderm bacteria have a thick PG layer on a single cell membrane and diderm bacteria feature two cell membranes, the inner and outer membranes, confining a periplasmic space where the PG layer is present. The PG layer of monoderm bacteria is exposed on the outer surface and hence can be directly observed. In contrast, it is difficult to observe the PG layer in diderm bacteria because it is covered by the outer membrane [13]. In this chapter, we describe the QFDE method to visualize the PG structures purified from E. coli cells.

2

Materials

2.1

Bacterial Strain

1. Escherichia coli DH5α.

2.2

Cell Culture

1. Luria-Bertani (LB) broth: 1% (w/v) Bacto tryptone, 0.5% (w/v) yeast extract, 1% (w/v) NaCl. 2. Glass test tube with an aluminum cap.

2.3

Cell Suspension

1. Phosphate-buffered saline (PBS): 75 mM sodium phosphate, pH 7.3, 68 mM NaCl. 2. 10% (w/v) Sodium dodecyl sulfate (SDS). 3. 10 mg/mL Chymotrypsin. 4. 1.5 mL microtube. 5. Micro tube lock (e.g., Greiner safety lock J616778). 6. Microcentrifuge. 7. Block heater at 96 °C.

2.4 Quick-Freezing (See Fig. 1)

1. Diamond compound #60000. 2. Filter paper. 3. Aluminum disk: 13 mm diameter with two square parts, 0.15 mm thick.

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Fig. 1 Quick-freezing machine. (a) Whole image of the freezing machine. (b) Magnified image of the freezing device. (c) Copper block unit. Top: copper block; middle: socket; Bottom: copper block set in the socket. (d) Aluminum disk set on the freezing base. (e) Schematic illustration of sample assembly on the aluminum disk. Number labels indicate: (1) freezing device, (2) liquid helium tank, (3) freezing base, (4) copper block assembly, (5) aluminum disk with sample, (6) freezing base, (7) sample and mica flake suspension, (8) rabbit lung tissue, (9) filter paper, (10) plastic spacer, and (11) aluminum disk

4. The plastic spacer: 13 mm diameter, 1.3 mm wide, and 1 mm thick. 5. Filter paper: 5 mm diameter. 6. Rabbit lung tissue: 3 mm2, 1 mm thick. 7. Mica flake [14]. 8. Tweezer. 9. Freezing machine (e.g., CryoPress). 10. Liquid helium. 11. Liquid nitrogen.

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Fig. 2 Replica machine. (a) Whole image of the replica machine (JEOL JFDV). (b) Magnified image of the sample chamber viewed from the front window. (c) Sample assembly in liquid nitrogen. (d) Cleaning of sample before fracture. (e) Sample on the stage just after fracture. Number labels indicate: (1) sample chamber window, (2) angle scale for sample stage, (3) sample stage, (4) knife, (5) thickness monitor for coating, (6) plastic spacer, (7) sample on the aluminum disk, (8) paint brush, and (9) fractured sample 2.5 Platinum Replica (See Fig. 2)

1. Freeze-etching device (e.g., JEOL JFDV). 2. Paint brushes. 3. Aluminum cap. 4. Polystyrene petri dish: 4 cm diameter. 5. 40% (w/v) Hydrogen fluoride (see Note 1). 6. Chlorine bleach for clothing. 7. 0.01% (w/v) SDS. 8. Copper 400 mesh grid for EM. 9. Glow discharge machine (e.g., Vacuum Device PIB-10). 10. Flatten-tip glass rod.

3 3.1

Methods Cell Culture

1. Inoculate a 20 μL volume of thawed frozen E. coli stock into a 5 mL volume of LB medium in a capped glass test tube. 2. Incubate overnight at 37 °C with shaking at 100 rpm until the culture growth reaches the stationary phase.

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SDS Treatment

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1. Centrifuge 5 mL of cell culture at 10,000 × g, 25 °C for 5 min. 3. Discard the supernatant and suspend the cell pellet in 1 mL of PBS. 4. Transfer the cell suspension to a 1.5 mL tube and centrifuge it at 10,000 × g at 25 °C for 5 min. 5. Discard the supernatant and suspend the cell pellet in 0.5 mL of PBS. 6. Add 0.5 mL of 10% (w/v) SDS. 7. Mix the cell suspension gently by inverting the tube. 8. Set a lock to the tube. 9. Incubate the tube at 96 °C for 3 h. 10. Incubate the tube at 25 °C for 20 min. 11. Centrifuge the tube at 20,000 × g at 25 °C for 30 min (see Note 2). 12. Discard the supernatant and suspend the cell pellet in 1 mL of PBS (see Note 3). 13. Centrifuge the tube at 20,000 × g at 25 °C for 30 min. 14. Discard the supernatant and suspend the cell pellet in 1 mL of PBS. 15. Add 10 μL of 10 mg/mL chymotrypsin to the cell suspension. 16. Place the tube in a block heater set at 37 °C. 17. Incubate the tube at 37 °C for 2 h. 18. Centrifuge the tube at 20,000 × g at 25 °C for 30 min. 19. Discard the supernatant and suspend the cell pellet in 1 mL of water. 20. Centrifuge the tube at 20,000 × g at 25 °C for 30 min. 21. Discard the supernatant and suspend the cell pellet in 1 mL of water. 22. Centrifuge the tube at 20,000 × g at 25 °C for 30 min. 23. Discard the supernatant and suspend the cell pellet in 50 μL of water.

3.3 Negative Staining EM

1. Glow-discharge carbon coated EM grids for 30 s. 2. Apply 5 μL of the sample solution onto the EM grid and incubate for 30 s. 3. Remove the extra solution with filter paper. 4. Apply 5 μL of 2% (w/v) phosphotungstic acid, pH 7.0 onto the EM grid and remove the solution quickly. 5. Dry the stained EM grid.

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Fig. 3 Negative-stained TEM image of isolated peptidoglycan layer of Escherichia coli. Boxed area in the left panel is magnified on the right

6. Observe the negative stained EM grid by TEM (e.g., JEM-1010, JEOL, Japan) operating at 80 kV, and capture the images with a camera (e.g., TemCam-F214 CCD camera, TVIPS, Germany) (Fig. 3) (see Note 4). 3.4

Quick-Freezing

1. To set samples on an aluminum disk, glue filter paper with a diameter of 5 mm and plastic spacer with epoxy adhesive and cyanoacrylate, respectively (Fig. 1d, e). 2. Set the aluminum disk on the base. 3. Place a rabbit lung slab on the aluminum disk (see Note 5). 4. Place 10 μL of mica flake solution on the lung slab. 5. Stir 10 μL of sample with mica flakes and absorb the extra sample. 6. Polish the surface of a copper block with a diamond compound (Fig. 1c). 7. Wash the copper block with pure water (see Note 6). 8. Set the copper block in the device and let the liquid helium flow at a gas rate of 5 L/min for 3 min to cool it (Fig. 1a). 9. Set the base with the aluminum disk on the freezing machine and freeze it (Fig. 1b). 10. Slide the aluminum disk from the base and transfer it to a Dewar bottle containing liquid nitrogen (see Note 7).

3.5 Platinum Shadowing and Carbon Backing

1. Pour liquid nitrogen into a device and evacuate to 10-5 Pa (Fig. 2a, b). 2. Remove plastic spacers from the sample on the aluminum disk in liquid nitrogen.

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3. Set the sample on the aluminum disk on the cooled replica base (Fig. 2c). 4. Remove frost from the sample surface using a pre-cooled brush (Fig. 2d). 5. Cover the sample with an aluminum cap. 6. Set the base onto the freeze-etching device (JOEL JFDV). 7. Set the temperature to -120 °C and wait for 10 min. 8. Remove the aluminum cap, heat it to -104 °C and wait for 3 min. 9. Fracture the surface with a knife cooled at -170 °C (Fig. 2e). 10. Move the knife to a position above the sample and wait for 10 min. 11. Coat the specimen with platinum to a thickness of 2 nm at an angle of 20° with 60 rpm rotation. 12. Coat carbon to a thickness of 10 nm at an angle of 80° with 60 rpm rotation. 3.6 Recovery and Observation of the Platinum Replica

1. Put 3 mL of 40% (w/v) hydrogen fluoride onto a petri dish and add few drops of 0.01% (w/v) SDS. 2. Gently float the replica with the lung slab on 40% (w/v) hydrogen fluoride (Fig. 4a). 3. Leave it for 1 h to detach the replica from the lung slab. 4. Pick up the replica with a flatten-tip glass rod and float it on fresh water for 5 min. 5. Pick up the replica with the flatten-tip glass rod and gently float it on a bleaching agent (Fig. 4b, c). 6. Leave it for 1 h. 7. Pick up the replica with the flatten-tip glass rod and float it on water for 5 min. 8. Pick up the replica with a glow-discharged copper grid and dry (Fig. 4d, e). 9. Observe the replica by TEM (Fig. 5).

4

Notes 1. Hydrofluoric acid is highly corrosive and must be used with proper precaution and protection. 2. Do not cool SDS treated sample while centrifuging. 3. PG pellets are transparent and difficult to see with the naked eye. 4. Confirm that all cells are lysed and that there is no damage to the PG sack.

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Fig. 4 Replica treatment. (a) Replica floating on hydrofluoric acid. (b) Picking up the replica by a flatten-tip glass rod. (c) Replica floating on the bleach reagent. (d) Replica floating on water. (e) Picking up replica by a copper grid. Number labels indicate: (1) flatten-tip glass rod, (2) plastic petri dish, (3) rabbit lung tissue, (4) replica, and (5) copper grid

Fig. 5 TEM image of replica of purified peptidoglycan layer of Escherichia coli. Boxed area in the left panel is magnified on the right

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5. The lung slab is used to absorb the mechanical shock for the sample and moisten it, and it dissolves itself easily during the replica pick-up. 6. Dry up the copper block completely. If water remains on them, they will stick upon freezing. 7. Frozen samples can be stored over many years in liquid nitrogen.

Acknowledgments We thank Junko Shiomi for technical support. This work was supported by Grants-in-Aid for Scientific Research (A) (MEXT KAKENHI, Grant Number JP17H01544), JST CREST (Grant Number JPMJCR19S5), and the Osaka City University (OCU) Strategic Research Grant 2019 to MM. References 1. Errington J (2013) L-form bacteria, cell walls and the origins of life. Open Biol 3(1):120143. https://doi.org/10.1098/rsob.120143 2. Turner RD, Vollmer W, Foster SJ (2014) Different walls for rods and balls: the diversity of peptidoglycan. Mol Microbiol 91(5):862–874. https://doi.org/10.1111/mmi.12513 3. Erickson HP (2017) How bacterial cell division might cheat turgor pressure – a unified mechanism of septal division in gram-positive and gram-negative bacteria. BioEssays 39(8). https://doi.org/10.1002/bies.201700045 4. Osawa M, Erickson HP (2018) Turgor pressure and possible constriction mechanisms in bacterial division. Front Microbiol 9:111. https://doi.org/10.3389/fmicb.2018.00111 5. Miyata M, Robinson RC, Uyeda TQP et al (2020) Tree of motility – a proposed history of motility systems in the tree of life. Genes Cells 25(1):6–21. https://doi.org/10.1111/ gtc.12737 6. Stulberg ER, Lozano GL, Morin JB et al (2016) Genomic and secondary metabolite analyses of streptomyces sp. 2AW provide insight into the evolution of the cycloheximide pathway. Front Microbiol 7:573. https://doi. org/10.3389/fmicb.2016.00573 7. Kunzler M (2018) How fungi defend themselves against microbial competitors and animal predators. PLoS Pathog 14(9):e1007184. https://doi.org/10.1371/journal.ppat. 1007184

8. McBride MJ, Nakane D (2015) Flavobacterium gliding motility and the type IX secretion system. Curr Opin Microbiol 28:72–77. https://doi.org/10.1016/j.mib.2015.07.016 9. Nan B, Zusman DR (2016) Novel mechanisms power bacterial gliding motility. Mol Microbiol 101(2):186–193. https://doi.org/10.1111/ mmi.13389 10. Heuser JE, Reese TS, Dennis MJ et al (1979) Synaptic vesicle exocytosis captured by quick freezing and correlated with quantal transmitter release. J Cell Biol 81(2):275–300. https:// doi.org/10.1083/jcb.81.2.275 11. Heuser JE (2011) The origins and evolution of freeze-etch electron microscopy. J Electron Microsc 60(Suppl 1):S3–S29. https://doi. org/10.1093/jmicro/dfr044 12. Megrian D, Taib N, Witwinowski J et al (2020) One or two membranes? Diderm firmicutes challenge the gram-positive/gram-negative divide. Mol Microbiol 113(3):659–671. https://doi.org/10.1111/mmi.14469 13. Tulum I, Tahara YO, Miyata M (2019) Peptidoglycan layer and disruption processes in Bacillus subtilis cells visualized using quickfreeze, deep-etch electron microscopy. Microscopy 68(6):441–449. https://doi.org/10. 1093/jmicro/dfz033 14. Heuser JE (1983) Procedure for freeze-drying molecules adsorbed to mica flakes. J Mol Biol 169(1):155–195. https://doi.org/10.1016/ s0022-2836(83)80179-x

Part VI Unique Motility System in Bacteria

Chapter 25 Purification and Structural Analysis of the Gliding Motility Machinery in Mycoplasma mobile Takuma Toyonaga and Makoto Miyata Abstract Isolating functional units from large insoluble protein complexes are a complex but valuable approach for quantitative and structural analysis. Mycoplasma mobile, a gliding bacterium, contains a large insoluble protein complex called gliding machinery. The machinery contains several chain structures formed by motors that are evolutionarily related to the F1-ATPase. Recently, we developed a method to purify functional motors and their chain structures using Triton X-100 and a high salt concentration buffer and resolved their structures using electron microscopy. In this chapter, we describe the processes of purification and structural analysis of functional motors for the gliding of M. mobile using negative-staining electron microscopy. Key words Mycoplasma gliding, Motor, F1-ATPase, Evolution, Chain structure, Solubilization, Electron microscopy, Single-particle analysis

1

Introduction Mycoplasmas are parasitic bacteria belonging to the class Mollicutes, which are known for their short genomes and lack of peptidoglycan layers [1–3]. Over ten mycoplasma species form a membrane protrusion at the cell pole and glide in the direction of the protrusion [4–6]. These motilities are thought to have evolved from the movement of cytoskeletal proteins and intracellular enzymes involved in housekeeping activities [3]. Mycoplasma mobile is a fish-pathogenic mycoplasma that adheres to solid surfaces covered with sialylated oligosaccharides and uses ATP as the energy source to glide at a speed of up to 4.5 μm/s (Fig. 1a) [5]. The gliding motility is thought to have evolved from the movement of F1-ATPase, based on sequence and structural analyses [7–9]. The machinery for the gliding motility can be divided into surface and internal structures (Fig. 1b). The surface structure has a leg protein for gliding, Gli349, which can bind to sialylated

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Fig. 1 Mycoplasma mobile gliding. Purple arrows indicate the gliding directions. (a) Phase-contrast microscopy of M. mobile. (Reproduced from Toyonaga et al. [9] with permission from the American Society for Microbiology). (b) Scheme of gliding machinery. The surface and internal structures are colored red and blue, respectively. Sialylated oligosaccharides on the host cell surface are depicted in orange

oligosaccharides [10–15]. The internal structure is a large complex consisting of a massive structure localized at the tip of the membrane protrusion and 28 chain-like structures lining the membrane [7, 8, 16]. Each chain is composed of an ATPase complex evolutionarily related to the F1-ATPase, suggesting that the chain generates the force for gliding. In this chapter, we refer to the chain as the motor chain and the ATPase complex as the motor. This large internal structure, including its motor chains, is challenging to isolate owing to its insoluble nature and propensity to form aggregates during centrifugation. Recently, we observed that the aggregated internal structure could be slightly disassembled by mechanical shearing in an NaCl-containing buffer and separated into motors and motor chains [9]. Thus, we successfully purified the motor, which features a hexameric structure like F1-ATPase and the motor chain. Here, we describe our protocol to purify the motor and motor chains and their structural analysis by negativestaining electron microscopy (EM). Briefly, M. mobile cells were treated with Triton X-100 and centrifuged to obtain internal structures. The internal structures were firmly pipetted into phosphate-buffered saline (PBS) supplemented with NaCl to solubilize the motors. The motors were subsequently purified by gel filtration. The motor chain was isolated by gentle treatment of the internal structures with high NaCl-containing buffers, followed by centrifugation. For structural analysis, each sample was stained with uranyl acetate solution on an EM grid, and particle images were acquired using an electron microscope. The particle images were then 2D classified to obtain the averaged images. Furthermore, the particle images of the motor chain selected by the 2D classification were used to reconstruct a 3D model.

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Materials Prepare all reagents with ultrapure water and analytical-grade reagents at 24–27 °C, unless otherwise stated.

2.1

Strains

1. Mycoplasma mobile mutant strain (P476R gli521) (see Note 1) [17].

2.2

Aluotto Medium

1. Dissolve 25.2 g of heart infusion broth and 6.72 g of yeast extract in 1080 mL of ultrapure water and adjust the pH to 7.8 by adding 0.42 mL of 10 M NaOH. 2. Autoclave at 121 °C for 20 min. 3. Cool down to 24–27 °C. 4. To this sterile media, add 120 mL of heat-inactivated horse serum, 1.2 mL of 50 mg/mL ampicillin solution, and 1.2 mL of 2.5 mg/mL amphotericin B solution. Perform this step on a clean bench.

2.3 Cell Growth and Harvest

1. 25 °C incubator. 2. 75 cm2 tissue culture flask. 3. 300 cm2 tissue culture flask. 4. Spectrophotometer.

2.4 Purification of Motor and Motor Chain

1. PBS: 8.1 mM Na2HPO4, 1.5 mM KH2PO4, pH 7.3, 2.7 mM KCl, and 137 mM NaCl (autoclave at 121 °C for 20 min). 2. Triton X-100 solution: 2% (w/v) Triton X-100, 0.2 mg/mL DNase, 10 mM MgCl2, and 2 mM phenylmethylsulfonyl fluoride in PBS. 3. Suspension buffer: PBS supplemented with 5 mM MgCl2. 4. Shaker. 5. Refrigerated centrifuge. 6. Sonicator. 7. 500 mL bottle. 8. 50 mL tube. 9. 15 mL tube. 10. 1.5 mL microtube. 11. Fast protein liquid chromatography system. 12. Gel filtration column (e.g., HiLoad™ 16/600 Superdex™ 200 pg, Cytiva). 13. 0.45 μm pore-size filter.

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2.5 Sodium Dodecyl SulfatePolyacrylamide Gel Electrophoresis (SDSPAGE)

1. Equipment for SDS-PAGE electrophoresis. 2. Running buffer: 25 mM Tris, 192 mM glycine, 0.1% SDS. 3. 4× SDS-loading buffer: 250 mM Tris–HCl, pH 6.8, 40% (w/v) glycerol, 10% SDS, 0.1% (w/v) bromophenol blue, and 20% (v/v) 2-mercaptoethanol. 4. Heating block set at 95 °C. 5. 12.5% (w/v) SDS-polyacrylamide gel.

2.6 Electron Microscopy

1. Carbon-coated copper 400 mesh grids. 2. Filter paper. 3. 2% (w/v) Uranyl acetate. 4. 80 kV transmission electron microscope. 5. A charge-coupled device (CCD) camera (e.g., FastScan-F214 (T) CCD camera, TVIPS).

2.7 Image Collection and Structural Analysis

3

1. Imaging software (e.g., EM menu, TVIPS). 2. RELION 3.0 [18]. 3. Gctf [19].

Methods

3.1 Cultivation of M. mobile

1. Inoculate 0.5 mL of thawed M. mobile stock into 20 mL of fresh Aluotto medium in a 75 cm2 tissue culture flask on a clean bench and incubate it at 25 °C for 2 days (see Note 2). 2. Inoculate 5 mL of culture into four 300 cm2 tissue culture flasks, each containing 300 mL of fresh Aluotto medium, on a clean bench and incubate at 25 °C for 2–3 days. 3. Measure OD600 of the culture after incubation using a spectrophotometer. Proceed to the next step when the cell density reaches an OD600 of 0.08.

3.2 Purification of Motor for Gliding

1. Transfer the cultures into 500 mL bottles. 2. Centrifuge the bottles (18,000 × g, 30 min, 4 °C). 3. Discard the supernatant and suspend the cell pellet in 30 mL of PBS. 4. Transfer the suspension to 50 mL tubes and centrifuge them (14,000 × g, 30 min, 4 °C). 5. Repeat steps 3 and 4 once. 6. Discard the supernatant and resuspend the pellet in 12.8 mL of PBS. 7. Transfer each 400 μL suspension to a 1.5 mL tube. 8. Sonicate the tubes for 1 min at 24–27 °C (see Note 3).

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9. Divide the treated solution evenly into eight 50 mL tubes. 10. Add PBS to each tube and adjust the volume to 12.5 mL. 11. Add an equal volume of Triton X-100 solution to each tube at 4 °C (see Note 4). 12. Shake the tubes gently for 1 h at 4 °C and centrifuge them (20,000 × g, 20 min, 4 °C). 13. Discard the supernatant and suspend the pellet in 40 mL of suspension buffer. 14. Centrifuge the tubes (20,000 × g, 20 min, 4 °C). 15. Discard the supernatant and resuspend the pellet in 5 mL of suspension buffer. 16. Transfer each 200 μL suspension to a 1.5 mL tube (see Note 5). 17. Resuspend the suspension in each tube by pipetting up and down ten times using a low-volume pipette (see Notes 6 and 7). 18. Incubate the samples overnight at 4 °C. 19. Repeat step 17 once. 20. Centrifuge the tubes (20,000 × g, 20 min, 4 °C). 21. Filter 5 mL of the supernatant using a 0.45 μm pore size filter and load it into a gel filtration column equilibrated with PBS and 1 mM MgCl2 (Fig. 2a) (see Notes 8 and 9). 22. Collect the peak fraction and confirm that the fraction contains the five motor components by SDS-PAGE and Coomassie Brilliant Blue (CBB) staining (Fig. 2b) (see Note 10).

Fig. 2 Purification of motor and motor chain. (a) The gel filtration elution pattern of the motor. The peak of the motor is marked by a black triangle. (b) Steps involved in the purification of the motor. Lane 1, cell lysate of M. mobile; lanes 2 and 3, detergent-soluble/detergent-insoluble fractions; lane 4, supernatant in step 20 of “Purification of motor for gliding”; lane 5, peak fraction of gel filtration. (c) Steps involved in the purification of the motor chain. Lane 1, detergent-insoluble fraction; lane 2, supernatant in step 15 of “Purification of motor chain for gliding.” Each fraction was subjected to SDS-PAGE and stained with CBB. Bands of motor components are marked by black triangles. Molecular masses are shown on the left. (Reproduced from Toyonaga et al. [9] with permission from the American Society for Microbiology)

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3.3 Purification of the Gliding Motor Chain

1. Transfer 15 mL of culture to 15 mL tubes. 2. Centrifuge the tubes (14,000 × g, 30 min, 4 °C). 3. Discard the supernatant and suspend the cell pellet in 1 mL of PBS. 4. Transfer the suspension to 1.5 mL tubes and centrifuge them (14,000 × g, 5 min, 4 °C). 5. Repeat steps 3 and 4 once. 6. Discard the supernatant and resuspend the pellet in 150 μL of PBS. 7. Sonicate the tube for 1 min at 24–27 °C. 8. Add an equal volume of Triton X-100 solution to the tube at 4 °C. 9. Shake gently at 4 °C for 30 min. 10. Centrifuge the tube (20,000 × g, 20 min, 4 °C). 11. Discard the supernatant and gently wash the surface of the pellet with 150 μL of the suspension buffer (see Note 11). 12. Suspend the pellet in 60 μL of suspension buffer. 13. Add 60 μL of suspension buffer containing 500 mM NaCl to the suspension and gently mix by pipetting the solution up and down twice (see Note 12). 14. Centrifuge the tube (5000 × g, 5 min, 4 °C). 15. Collect the supernatant. 16. Analyze the samples by SDS-PAGE and CBB staining, if required (Fig. 2c).

3.4 Specimen Preparation for Negative-Staining EM and Data Collection

1. Glow-discharge the carbon-coated copper grids on a filter paper for 1 min. 2. Place 4 μL of sample solution on the grid and incubate at 24–27 °C for 1 min. 3. Remove the solution from the grid with filter paper, add 4 μL of 2% (w/v) uranyl acetate solution for staining, and incubate at 24–27 °C for 30 s. 4. Remove the solution from the grid with filter paper and allow the grid to dry on the filter paper for at least 30 min. 5. Observe the grid using a transmission electron microscope equipped with a CCD camera. 6. Acquire images at a nominal magnification of 40,000×, corresponding to 2.58 Å/pixel, with a dose rate of 100 e-/ Å2/s and an exposure time of 1 s (Fig. 3).

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Fig. 3 EM images of (a) motor and (b) motor chain. (Top) Electron micrograph of negatively stained particles. (Bottom) Representative 2D averaged images. The motor chain structures are marked by blue lines. (Reproduced from Toyonaga et al. [9] with permission from the American Society for Microbiology) 3.5 Image Processing of the Motor Proteins

1. Import the field images of motors into RELION 3.0. 2. Estimate the parameters for the contrast transfer function using the Gctf program. 3. Manually pick up over 150 motor particles and extract them with a box size of 180 × 180 pixels. 4. Bin the particles to 5.16 Å/pixel and 2D-classify them into four to eight classes. 5. Over 150 motor particles are automatically picked up using three to four 2D average images obtained as templates and subsequently extracted. 6. Bin the particles to 5.16 Å/pixel and 2D-classify them into 100 classes.

3.6 Image Processing of the Motor Chain

1. Import the field images of motor chains into RELION 3.0. 2. Estimate the parameters for the contrast transfer function using the Gctf program. 3. Manually pick approximately 2000 particles for the motor chain and extract them with a box size of 276 × 276 pixels with 50% overlap (see Note 13). 4. Bin the particles to 5.16 Å/pixel and 2D classify them into 20 classes. 5. Select all optimal, averaged images and 2D classify the particles used into 20 classes.

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6. Select all optimal, averaged images and create a 3D model from the used particles using the “initial model” with initial and final resolution limits of 100 Å and 50 Å, respectively. 7. Refine the model using “3D auto-refine” to obtain the final 3D density map.

4

Notes 1. The wild-type M. mobile strain 163 K (ATCC 43663) can also be used. 2. The depth of the solution should not exceed 20 mm. 3. Sonication is required for dispersing cells into solution. 4. Adding Triton X-100 brings the total volume of the solution to 200 mL. The purpose of large-volume treatment is to dissolve as many mycoplasma cells as possible, leaving no potentially soluble components in the pellet after the subsequent centrifugation step. 5. A smaller-volume treatment is advantageous for suspending the pellet. 6. Steps 16–18 are required for motor solubilization. 7. If the suspension buffer is replaced by a buffer with a higher salt concentration (e.g., buffer: 20 mM Tris–HCl, pH 7.5, 350 mM NaCl), the solubilization efficiency will increase. However, the final purity will be reduced owing to the contamination of other solubilized proteins. Additionally, motor aggregation likely occurs when the final salt concentration is reduced. 8. The motor fraction was recovered as a peak at the void volume in the HiLoad™ 16/600 Superdex™ 200 pg column. 9. To obtain a motor fraction with a single hexamer, treat the supernatant with 1.5% (w/v) sodium cholate overnight before loading it onto the column. 10. If the motor concentration is low, concentrate the motor using an ultrafiltration unit. Add approximately 0.1% CHAPS to the solution to prevent protein adsorption to the filter membrane, if necessary. 11. Repeating steps 10–12 may result in a better purity of the motor chain fraction. 12. After step 13, the chain structures dissociate over time and may be lost during the process. Therefore, the subsequent steps should be performed as quickly as possible. 13. The connecting parts of the two motors are centered in the box to pick up the particles.

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Acknowledgments This study was supported by a grant-in-aid for scientific research on the Innovative Area Harmonized Supramolecular Motility Machinery and Its Diversity (MEXT KAKENHI grant number JP24117002), by grants-in-aid for scientific research (A) (MEXT KAKENHI grant number JP17H01544), by JST CREST grant number JPMJCR19S5, Japan, by the Osaka City University (OCU) Strategic Research Grant 2018 for top priority researches to M.M. References 1. Razin S, Hayflick L (2010) Highlights of mycoplasma research--an historical perspective. Biologicals 38(2):183–190 2. Grosjean H, Breton M, Sirand-Pugnet P et al (2014) Predicting the minimal translation apparatus: lessons from the reductive evolution of mollicutes. PLoS Genet 10(5):e1004363 3. Miyata M, Robinson RC, Uyeda TQP et al (2020) Tree of motility – a proposed history of motility systems in the tree of life. Genes Cells 25(1):6–21 4. Miyata M (2010) Unique centipede mechanism of Mycoplasma gliding. Annu Rev Microbiol 64:519–537 5. Miyata M, Hamaguchi T (2016) Prospects for the gliding mechanism of Mycoplasma mobile. Curr Opin Microbiol 29:15–21 6. Miyata M, Hamaguchi T (2016) Integrated information and prospects for gliding mechanism of the pathogenic bacterium Mycoplasma pneumoniae. Front Microbiol 7:960 7. Nakane D, Miyata M (2007) Cytoskeletal “jellyfish” structure of Mycoplasma mobile. Proc Natl Acad Sci U S A 104(49):19518–19523 8. Nishikawa M, Nakane D, Toyonaga T et al (2019) Refined mechanism of Mycoplasma mobile gliding based on structure, ATPase activity, and sialic acid binding of machinery. mBio 10(6):e02846–e02819 9. Toyonaga T, Kato T, Kawamoto A et al (2021) Chained structure of dimeric F(1)-like ATPase in Mycoplasma mobile gliding machinery. mBio 12(4):e0141421 10. Uenoyama A, Kusumoto A, Miyata M (2004) Identification of a 349-kilodalton protein (Gli349) responsible for cytadherence and glass binding during gliding of Mycoplasma mobile. J Bacteriol 186(5):1537–1545

11. Metsugi S, Uenoyama A, Adan-Kubo J et al (2005) Sequence analysis of the gliding protein Gli349 in Mycoplasma mobile. Biophysics (Nagoya-shi) 1:33–43 12. Nagai R, Miyata M (2006) Gliding motility of Mycoplasma mobile can occur by repeated binding to N-acetylneuraminyllactose (sialyllactose) fixed on solid surfaces. J Bacteriol 188(18): 6469–6475 13. Kasai T, Hamaguchi T, Miyata M (2015) Gliding motility of Mycoplasma mobile on uniform oligosaccharides. J Bacteriol 197(18): 2952–2957 14. Mizutani M, Tulum I, Kinosita Y et al (2018) Detailed analyses of stall force generation in Mycoplasma mobile gliding. Biophys J 114(6): 1411–1419 15. Hamaguchi T, Kawakami M, Furukawa H et al (2019) Identification of novel protein domain for sialyloligosaccharide binding essential to Mycoplasma mobile gliding. FEMS Microbiol Lett 366(3):fnz016 16. Kobayashi K, Kodera N, Kasai T et al (2021) Movements of Mycoplasma mobile gliding machinery detected by high-speed atomic force microscopy. mBio 12(3):e0004021 17. Uenoyama A, Seto S, Nakane D et al (2009) Regions on Gli349 and Gli521 protein molecules directly involved in movements of Mycoplasma mobile gliding machinery, suggested by use of inhibitory antibodies and mutants. J Bacteriol 191(6):1982–1985 18. Zivanov J, Nakane T, Forsberg BO et al (2018) New tools for automated high-resolution cryoEM structure determination in RELION-3. elife 7:e42166 19. Zhang K (2016) Gctf: real-time CTF determination and correction. J Struct Biol 193(1): 1–12

Chapter 26 Motility Assays of Mycoplasma mobile Under Light Microscopy Taishi Kasai and Makoto Miyata Abstract Mycoplasma mobile forms a membrane protrusion at a pole as an organelle. M. mobile cells bind to solid surfaces and glide in the direction of the protrusion. In gliding motility, M. mobile cells catch, pull and release sialylated oligosaccharides on host cells. The observation of Mycoplasma species under light microscopy is useful for the analysis of adhesion ability and the motility mechanism. Key words Mycoplasma, Bacterial motility, Gliding motility, Tunnel chamber, Phase-contrast microscope

1

Introduction Mollicutes are wall-less gram-positive bacteria [1–3]. Mollicutes are classified into four subgroups, Hominis, Pneumoniae, Spiroplasma, and Phytoplasma [4, 5]. Mycoplasma mobile, which is classified to Hominis subgroup, forms a membrane protrusion at a cell pole and glides on solid surfaces [6]. The gliding motility system of M. mobile is not related to any other bacterial motility systems, such as flagella and pili [3, 7]. M. mobile is the fastest in the gliding species. The gliding speed ranges from 2.0 to 4.5 μm per second, or three to seven times the length of the cell body per second [6, 8]. The machinery of gliding motility is composed of three large proteins and cytoskeletal structures [6, 9, 10]. One of the three proteins, called Gli349, acts as a “leg” for the gliding motility. The Gli349 protein catches, pulls, and releases sialylated oligosaccharides on host cell surfaces [11–18]. Here, I describe how to observe the adhesion and motility of M. mobile under light microscopy.

Tohru Minamino et al. (eds.), Bacterial and Archaeal Motility, Methods in Molecular Biology, vol. 2646, https://doi.org/10.1007/978-1-0716-3060-0_26, © The Author(s), under exclusive license to Springer Science+Business Media, LLC, part of Springer Nature 2023

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Materials Cultured Cell

1. Aluotto medium: 2.1% (w/v) heart infusion broth, 0.56% (w/v) yeast extract, 10% (v/v) horse serum, 25 μg/mL amphotericin B, 50 μg/mL ampicillin (see Note 1). Store at 4 °C. 2. Mycoplasma mobile 163K, ATCC43663. 3. 100 mL Erlenmeyer flask (see Note 2). 4. Incubator adjusted at 25 °C. 5. 1.5 mL microcentrifuge tube. 6. Centrifuge.

2.2

Tunnel Chamber

1. Slide glass. 2. Cover slip. 3. Double-sided tape. 4. Filter paper. 5. Humidity chamber. 6. Vaseline.

2.3 Microscopic Observation and Image Analysis

1. Phase-contrast microscope. 2. Plan Apochromat 100× oil immersion objective lens. 3. CCD camera. 4. ImageJ (http://imagej.nih.gov/ij/).

3

Methods

3.1 Preparation of Tunnel Chambers

1. Adhere two pieces of double-sided tapes 5 mm apart on glass surface. 2. Remove the release paper and place a cover slip on the doublesided tapes. 3. Press the cover slip to achieve tight binding (Fig. 1a).

3.2 Preparation of Cells

1. Inoculate 10 mL Aluotto medium in a 100 mL Erlenmeyer flask with 1 mL frozen stock of Mycoplasma mobile. 2. Incubate the culture for 13–16 h at 25 °C until OD achieves 0.06–0.08 at a wavelength of 600 nm. 3. Collect 1 mL overnight culture to a microcentrifuge tube. 4. Centrifuge the tube at 12,000 g for 4 min at 25 °C. 5. Discard the supernatant by a pipet. 6. Resuspend the cell pellet in 100 μL fresh Aluotto medium.

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Fig. 1 Tunnel chamber assembly for motility assay of M. mobile. (a) A cover slip is combined to glass slide with double-sided tape. (b) Injection of cell suspension and buffer to the chamber. The black arrows show the flow of solution. (c) Sealing of two chamber ports by vaseline

Fig. 2 Phase-contrast microscopic images of M. mobile cells observed in tunnel chamber at different stages. The cell solution is injected into the tunnel chamber at time zero. Scale bar, 10 μm 3.3 Observation and Analysis of Mycoplasma Motility

1. Insert 20 μL cell suspension into a tunnel chamber. Absorb the overflowing suspension by a piece of filter paper (Fig. 1b). 2. Place the tunnel chamber with a cover slip side facing downward and incubate it for 10 min at 25 °C in a humidity chamber (Fig. 2). 3. Insert 50 μL fresh Aluotto medium twice to remove floating cells (Fig. 2).

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4. Seal both ports of the tunnel chamber with vaseline (Fig. 1c). 5. Set the tunnel chamber onto a stage of a phase-contrast microscope equipped with a 100 × oil immersion objective lens. 6. Record microscopic images with a CCD camera (see Note 3). 7. Select the binding cells using ge>Adjust>Threshold” of ImageJ.

a

command

“Ima-

8. Measure the XY coordinates of cells for all frames using a command “Analyze> Analyze particles.” 9. Calculate the gliding speed from the displacement of the cells between each frame.

4

Notes 1. Autoclave the solution of heart infusion broth and yeast extract. Horse serum is heated to 56 °C for 30 min. The heat treatment is the inactivation of complement activity in the horse serum. Cool down the medium to 56 °C and add the heat-inactivated horse serum and antibiotics. 2. A 25 cm2 tissue culture flask may be used instead of an Erlenmeyer flask. 3. The frame rate of CCD camera is higher than 30 frames per second to accurately track cells.

References 1. Razin S, Hayflick L (2010) Highlights of mycoplasma research—an historical perspective. Biologicals 38:183–190. https://doi. org/10.1016/j.biologicals.2009.11.008 2. Grosjean H, Breton M, Sirand-Pugnet P et al (2014) Predicting the minimal translation apparatus: lessons from the reductive evolution of mollicutes. PLoS Genet 10:e1004363. https://doi.org/10.1371/journal.pgen. 1004363 3. Miyata M, Robinson RC, Uyeda TQP et al (2020) Tree of motility – a proposed history of motility systems in the tree of life. Genes Cells 25:6–21. https://doi.org/10.1111/gtc. 12737 4. Barre´ A, de Daruvar A, Blanchard A (2004) MolliGen, a database dedicated to the comparative genomics of Mollicutes. Nucleic Acids Res 32:D307–D310. https://doi.org/10. 1093/nar/gkh114 5. Weisburg WG, Tully JG, Rose DL et al (1989) A phylogenetic analysis of the mycoplasmas: basis for their classification. J Bacteriol 171:

6455–6467. https://doi.org/10.1128/jb. 171.12.6455-6467.1989 6. Miyata M, Hamaguchi T (2016) Prospects for the gliding mechanism of Mycoplasma mobile. Curr Opin Microbiol 29:15–21. https://doi. org/10.1016/j.mib.2015.08.010 7. Nakamura S, Minamino T (2019) Flagelladriven motility of bacteria. Biomolecules 9: E 2 7 9 . h t t p s : // d o i . o r g / 1 0 . 3 3 9 0 / biom9070279 8. Miyata M (2010) Unique centipede mechanism of Mycoplasma gliding. Annu Rev Microbiol 64:519–537. https://doi.org/10.1146/ annurev.micro.112408.134116 9. Nishikawa MS, Nakane D, Toyonaga T et al (2019) Refined mechanism of Mycoplasma mobile gliding based on structure, ATPase activity, and sialic acid binding of machinery. mBio 10:e02846–e02819. https://doi.org/ 10.1128/mBio.02846-19 10. Tulum I, Kimura K, Miyata M (2020) Identification and sequence analyses of the gliding

Gliding Motility Assay machinery proteins from Mycoplasma mobile. Sci Rep 10:3792. https://doi.org/10.1038/ s41598-020-60535-z 11. Adan-Kubo J, Uenoyama A, Arata T, Miyata M (2006) Morphology of isolated Gli349, a leg protein responsible for Mycoplasma mobile gliding via glass binding, revealed by rotary shadowing electron microscopy. J Bacteriol 188:2821–2828. https://doi.org/10.1128/ JB.188.8.2821-2828.2006 12. Hamaguchi T, Kawakami M, Furukawa H, Miyata M (2019) Identification of novel protein domain for sialyloligosaccharide binding essential to Mycoplasma mobile gliding. FEMS Microbiol Lett 366:fnz016. https:// doi.org/10.1093/femsle/fnz016 13. Kasai T, Hamaguchi T, Miyata M (2015) Gliding motility of Mycoplasma mobile on uniform oligosaccharides. J Bacteriol 197:2952–2957. https://doi.org/10.1128/JB.00335-15 14. Kasai T, Nakane D, Ishida H et al (2013) Role of binding in Mycoplasma mobile and Mycoplasma pneumoniae gliding analyzed through inhibition by synthesized sialylated

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compounds. J Bacteriol 195:429–435. https://doi.org/10.1128/JB.01141-12 15. Lesoil C, Nonaka T, Sekiguchi H et al (2010) Molecular shape and binding force of Mycoplasma mobile’s leg protein Gli349 revealed by an AFM study. Biochem Biophys Res Commun 391:1312–1317. https://doi.org/10.1016/j. bbrc.2009.12.023 16. Metsugi S, Uenoyama A, Adan-Kubo J et al (2005) Sequence analysis of the gliding protein Gli349 in Mycoplasma mobile. Biophysics (Nagoya-shi) 1:33–43. https://doi.org/10. 2142/biophysics.1.33 17. Morio H, Kasai T, Miyata M (2016) Gliding direction of Mycoplasma mobile. J Bacteriol 198:283–290. https://doi.org/10.1128/JB. 00499-15 18. Nagai R, Miyata M (2006) Gliding motility of Mycoplasma mobile can occur by repeated binding to N-acetylneuraminyllactose (sialyllactose) fixed on solid surfaces. J Bacteriol 188:6469–6475. https://doi.org/10.1128/ JB.00754-06

Chapter 27 Detection of Steps and Rotation in the Gliding Motility of Mycoplasma mobile Yoshiaki Kinosita, Mitsuhiro Sugawa, Makoto Miyata, and Takayuki Nishizaka Abstract Mycoplasma mobile is one of the fastest gliding bacteria, gliding with a speed of 4.5 μm s
1. This gliding motility is driven by a concerted movement of 450 supramolecular motor units composed of three proteins, Gli123, Gli349, and Gli521, in the gliding motility machinery. With general experimental setups, it is difficult to obtain the information on how each motor unit works. This chapter describes strategies to decrease the number of active motor units to extract stepwise cell movements driven by a minimum number of motor units. We also describe an unforeseen motility mode in which the leg motions convert the gliding motion into rotary motion, which enables us to characterize the motor torque and energy-conversion efficiency by adding some more assumptions. Key words Bacteria, Mycoplasma mobile, Motility, Gliding, Optical microscope, Single-molecular techniques, Sialyllactose, Stepwise movement, Rotation

1

Introduction Bacteria show various types of motilities, such as gliding, swimming, and twitching, driven by surface appendages composed of supramolecular motility machinery [1]. Mycoplasma mobile is one of the fastest gliding bacteria with a speed of 4.5 μm s
1. The cell body is flask-shaped with a size of 1 μm and glides on solid surfaces in the direction of its protrusion [2]. Neither flagella nor pili are involved in this gliding movement [3]. Three proteins have been identified as surface proteins for Mycoplasma gliding: Gli123 as a scaffold for the assembly of other motility components [4], Gli349 as a leg repeating the binding to and releasing from sialylated oligosaccharides [5–8], Gli521 as a gear transmitting the intracellular movement to Gli349 (Fig. 1) [8–11]. The motor units localize around the cell neck, and the number of units has been estimated to be ~450. The intracellular structure of the motor unit comprises

Tohru Minamino et al. (eds.), Bacterial and Archaeal Motility, Methods in Molecular Biology, vol. 2646, https://doi.org/10.1007/978-1-0716-3060-0_27, © The Author(s), under exclusive license to Springer Science+Business Media, LLC, part of Springer Nature 2023

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Fig. 1 Schematics of the experimental setup. The motility of M. mobile was observed by an inverted microscope. M. mobile glides on a glass surface in the direction of the cell protrusion, and this motility is driven by a supramolecular motility machinery, which can be divided into the cell surface and internal structures. The cell surface structure is composed of three large proteins with different roles: Gli123 as a scaffold for other motility machinery, Gli349 as a leg binding to and releasing from sialylated oligosaccharides, and Gli521 as a gear transmitting the intracellular movement to Gli349. The internal structure consists of least ten proteins, and two of them have the walker A and Walker B motif for ATP binding and hydrolysis, respectively. Their amino acid sequences have similarity to α- and β-subunits of F-type ATPase. The change from cells to ghosts was judged by the reduction of image density by phase-contrast microscopy in real time. To trace M. mobile ghost movement with high spatiotemporal resolution, fluorescently-labeled ghosts were illuminated by a mercury lamp. The images captured by an EMCCD camera were analyzed by 2-D Gaussian fitting, and the positions were localized with a 2 nm accuracy. Em: emission filter, Ex: excitation filter, and DM: dichroic mirror

proteins homologous to the α-and β-subunits of F-type ATPase [12–15]. These data suggest that the F-type ATPase might be an ancestor of the motor for the Mycoplasma gliding motility, and in fact, a membrane-permeabilized ghost model has shown ATP-driven gliding movement [16]. The detection of stepwise motion is one of the most important experiments in the study of molecular motors [17]. The step size correlates with the periodicity of cytoskeleton and motor structures, and the waiting time between steps (dwell time) reflects the chemical and transition states such as ATP binding and substrate release. Therefore, the detection of steps reveals how the motor works at the molecular level, including the chemo-mechanical

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coupling and energy-conversion efficiency. In eukaryotes, reconstructions of motor assay systems by purified motor proteins enable easy access to this demand through bead attachment or fluorescent dye binding at a single-molecular level [17]. On the other hand, the reconstitution of motor machineries in prokaryotes has never been succeeded. Therefore, in the motor study of prokaryote, motor movement must be observed in situ. In addition to this restriction, the single-cell gliding movement of Mycoplasma is driven by multiple legs, making it very difficult to detect steps driven by each gliding unit. In this chapter, we describe methods to extract the unitary steps by adding free sialyllactose in solution to make most of the active legs bind to free sialyllactose, so that consequently, the cell motion by a minimum number of gliding units would be detected. We also described an unforeseen motility mode in which the leg motions convert the gliding motion into rotary motion [18]. This mode is seen when the ghost is prepared with a high concentration of detergent, where the ghost may be bound to the glass surface with a further reduced number of active legs for movement. These features allow us to observe a rotary motion as illustrated with a pivoting model and to characterize the energyconversion efficiency.

2

Materials All buffers are prepared with an ultrapure water (Milli-Q). All chemicals are purchased from commercial suppliers.

2.1

Strains

1. Mycoplasma mobile ATCC 43663 (WT strain for M. mobile). 2. M. mobile P476R gli521 mutant (This strain is recommended, because the proportion of cells attached to a glass surface after detergent treatment is higher compared to WT) [16].

2.2

Chemicals

1. Bovine serum albumin (BSA). 2. Triton X-100. 3. Methylcellulose (e.g., M0512; Sigma-Aldrich). 4. Cy3-NHS-ester (e.g., PA23001; GE Health care). 5. DNase. 6. N-acetylneuraminyllactose (e.g., Sialyllactose, A0828; SigmaAldrich) (see Note 1). 7. Dithiothreitol (DTT). 8. Adenosine triphosphate (ATP).

2.3

Stock Solution

1. Aluotto medium: 2.1% (w/v) heart infusion broth, 0.56% (w/v) yeast extract, 10% (v/v) horse serum, 0.005% (w/v) ampicillin.

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2. Buffer A: 75 mM sodium phosphate, pH 7.4, 68.4 mM NaCl, 50 mM glucose. 3. 1 M DTT. 4. 0.5 M EGTA pH 7.5 adjusted by 4 N NaOH. 5. Buffer B: 10 mM Tris–HCl, pH 7.5, 50 mM NaCl, 1 mM DTT, 1 mM EGTA, 2 mM MgCl2, 0.5 mg/mL BSA. 6. Buffer C: Buffer B containing 0.01% (w/v) Triton X-100. 7. 1% (w/v) Methylcellulose (see Note 2). 8. Buffer D: Buffer B containing 0.1% (w/v) methylcellulose and 1 mg/mL DNase. 9. 200 mM ATP pH 7.0 adjusted by 4 N NaOH. 10. Buffer E: buffer B containing ATP and 0.1% (w/v) methylcellulose. 11. 10 mM sialyllactose. 12. Buffer F: buffer D containing 1 mM sialyllactose. 13. Buffer G: Buffer B containing 0.013% (w/v) Triton X-100. 2.4 Fluorescence Microscopy (Fig. 1)

1. Inverted Microscope (e.g., Ti-E; Nikon Instruments) equipped with a 100 objective lens (e.g., Fluor 100 with Ph and 1.3 N.A.; Nikon Instruments) and a mercury lamp. 2. Optical table (e.g., RS-2000; Newport). A microscope was fixed on the table (see Note 3). 3. Filter set (e.g., TxRed4040B; Semrock): Dichroic mirror, emission filter, and excitation filter. 4. Electron multiplying charged coupled device camera (e.g., EMCCD camera, Ixon+ DU860; Andor technology).

2.5 Phase-Contrast Microscopy

1. Inverted microscope (e.g., IX71; Olympus) equipped with a 100 objective lens (e.g., UPLSAPO 100 with Ph and 1.4 N. A.; Olympus) and a halogen lamp. 2. Optical table (e.g., RS-2000; Newport). 3. High-speed scientific complementary metal oxide semiconductor camera (e.g., sCMOS camera, LRH1540N; Digimo).

2.6

Tunnel Slide

1. Double-sided tape. 2. 22  32 mm glass. 3. 18  18 mm glass.

2.7

Software

1. Image J 1.48v (http://rsb.info.nih.gov/ij/). 2. Andor solis software (Andor technology). 3. Igor pro. software 8.04 (Hulinks).

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Methods All experiments were performed at room temperature, unless indicated otherwise. All solutions in the tunnel slide were replaced on the microscope stage. A piece of filter paper was put on the other side to replace the solution in the tunnel slide.

3.1

Cultivation

1. Add 10 mL of Aluotto medium into a tissue culture flask. 2. Add 100 μL of frozen stock into medium. 3. Incubate the cells at 25 reaches 0.06.

3.2 Preparation of Fluorescently Labeled Cells



C for 2–3 days until OD600

1. Centrifuge 1 mL of cultured cells at 12,000  g for 4 min. 2. Discard supernatant by a pipette and resuspend the pellet in 1 mL buffer A containing 1 mg/mL of Cy3-NHS-ester. 3. Incubate for 2 h. 4. Centrifuge at 12,000  g for 4 min to remove excess Cy3 dye. 5. Discard supernatant and resuspend the pellet in 1 mL buffer A. 6. Centrifuge to remove excess Cy3-NHS-ester at 12,000  g for 4 min. 7. Discard supernatant and resuspend the pellet in 1 mL buffer A.

3.3 Construction of Tunnel Slide

1. Cut two pieces of ~30 mm length of a double-sided tape. 2. Fix two tapes onto 24  36 mm glass with an 5 mm intervals. 3. Place coverslip (18  18 mm) onto double-sided tapes.

3.4 Detection of Stepwise Movements

1. Infuse the Aluotto medium into a tunnel slide. 2. After 5 min, fluorescent-labeled cells were infused into the same chamber. 3. Wait for 10 min. 4. Remove unbound cells by infusing buffer A into the tunnel slide. 5. For the preparation of gliding ghosts, infuse the buffer C with observing the cells. 6. Infuse buffer D into the tunnel slide immediately after the image density of the cells decrease (see Note 4). 7. Subsequently, infuse buffer E. 8. Confirm the reactivation of ghost gliding and infuse buffer F (see Note 5). 9. Under mercury lamp excitation, fluorescently labeled ghosts are captured with an EMCCD camera for 30 s with a temporal resolution of 2 ms.

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Fig. 2 Stepwise gliding and rotary motion of M. mobile. (a) Strategies to analyze stepwise motion in ghosts gliding. A typical example of the x–y trace of gliding is shown in the upper left inset, where the ghost position was localized by 2-D Gaussian fitting. Black points in this trace were further analyzed as shown in the displacement graph. The arrow in the inset indicates the forward direction of gliding. The time course of ghost positions is presented with open circles (unfiltered data points), which was analyzed with the pairwise distance function (PDF) as shown in the lower inset. Red lines on the original data points indicate the lines fitted with the step-finding algorithm. The histogram obtained by the PDF analysis is shown in the lower inset. The vertical lines indicate the positions of 9 gaussian functions fitted to individual peaks of the histogram, as ΣiAiexp[
(x
iΔ)2/(2σ i2)] + A0exp[
(x
x0)2/(2σ 02)], where Δ is the unit step size and i ¼ 1–9. (b) Sequential images of a rotary ghost in CCW- (top) and CW-directional rotation (bottom) at 100 ms intervals. Scale bar,1 μm. (c) Traces of ghost rotation. The trajectories of the center of mass (left) and the integrated revolutions (right) are shown for CCW (blue) and CW (green). (The figures are reused with permission from Kinosita et al. [18] for (a) and Kinosita et al. [28], Springer Nature” for (b–d), with modification)

10. Save the sequential tif-images in 14-bit format without compression and use the image set for further analyses. 11. Perform 2-D Gaussian fitting on the image set using Igor Pro. software to localize the center of the image density of the gliding ghosts as x–y coordinates (Fig. 2a inset) (see Note 6) [19]. 12. Find out a section where a ghost has moved in a straight line (either x- or y-axis), excise the section as a ROI (region of interests), and adjust its angle for the following analysis. 13. Quantify the periodicity of the displacement by applying the pairwise distance function to the time course of displacement (see Note 7). 14. Fit the data with multiple Gaussian functions and estimate the step size from the periodicity of the peaks (Fig. 2a inset).

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15. Apply Kerssemakers analysis to the data showing four or more periodic steps in pairwise analysis to quantify the dwell time, which provides us information of chemical states, such as ATP binding and ATP hydrolysis (Fig. 2a) (see Note 8) [20]. 3.5 Detection of Rotation

1. Confirm gliding cells in a tunnel slide by a phase-contrast microscopy. 2. For the preparation of rotary ghosts, infuse the buffer G with observing the cells. 3. Infuse buffer D into the tunnel slide immediately after the optical density of the cells decreases (see Note 9). 4. Subsequently, infuse buffer E. 5. (Optional) Infuse buffer F to observe the stepwise rotation (see Note 10). 6. Record the sequential images for 30 s with a temporal resolution of 4 ms using a sCMOS camera and use the image set for further analyses (Fig. 2b). 7. Perform centroid fitting using Igor Pro. software to determine the centroid position of ghosts (Fig. 2c) (see Note 11). 8. Convert the x–y position of each frame into the angle (θ) following equation: θ ¼ arctan(y/x). 9. The time course of rotation is produced by dividing the integrated angle by 360 (Fig. 2d). 10. Fit the data linearly to quantify rotary speeds. 11. Calculate the generated torque by each ghost following equation: T ¼ 2πfξ, where f is rotational speed and ξ ¼ 8πηa3 + 6πηar2 the viscous drag coefficient, with r the radius of rotation of the ghost center, a the radius of ghost, and η the viscosity [21]. 12. If stepwise rotation is detected, apply the methods in Subheading 3.1, steps 11–13 to quantify the step size and Dwell time.

4

Notes 1. We used sialyllactose (A0828, Sigma-Aldrich) in our original paper [18]. However, we are not able to use this compound any more due to out of production. Sialyllactose is a mixture of 30 -sialyllactose (e.g., A8681: Sigma-Aldrich) and 60 -silalylactose (e.g., A8556: Sigma-Aldrich). 30 -sialyllactose might cause strong inhibition on M. mobile gliding [6]. 2. Methylcellulose is firstly warmed with stirring by stirrer in a beaker. When the powder disperses, the beaker is moved on to the ice. Overnight incubation is recommended to dissolve the powder completely.

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3. The microscope should be on the optical table with a vibration isolator, which reduces vibrations caused by people walking around, etc. This is very critical to extract nano-meter size movement driven by a tiny molecular motor, for example, 5.5 nm in Myosin II [22] and 8 nm in Kinesin I [23]. 4. Motility is driven by the chemical energy of ion flow or ATP hydrolysis, and therefore, the energy-conversion mechanism should be clarified. The ghost technique provides fruitful information because the cell membrane of gliding ghosts is permeabilized with detergent, enabling us to control the intracellular molecular function coupled with ATP hydrolysis. For the preparation of gliding ghosts, the change from live cells to ghosts by adding a detergent can be observed in real time by phase contrast microscopy. When the cell membrane is permeabilized with detergent, the cytoplasm is expelled, resulting in lower density images compared to those of live cells. 5. To prepare gliding ghosts, you should control the detergent concentration so that the image density of cells is immediately reduced to the level of the ghosts after addition of detergent. The image density of all live cells should instantly drop due to permeabilization of cell membrane. Under such a condition, roughly 70% of the ghosts will be reactivated. 6. The centroid position of the ghost (x, y) is determined by fitting each image with the following 2D Gaussian function with the cross-correlation term, cor: I0+ Aexp[
(x
xc)2/ 2σ x2 + (y
yc)2/2σ y2
2cor(x
xc)(y
yc)/σ xσ y]/(2 (1
cor2)), where I0 is the background intensity and σ x and σ y are the intensity variance. 7. Pairwise analysis has been used to extract the stepwise movements of molecular motors such as kinesin [24]. Pairwise analysis is a method in which the first frame (n) is fixed, and the difference in position of the next frame is calculated. This procedure is performed by the last frame, and then, this routine is again performed starting from n + 1 frames. Finally, we summarize the difference in a histogram. We applied this analysis to M. mobile for the quantification of its step sizes. When more than four periodic peaks in the histogram were seen, we applied the Kerssemakers analysis to quantify the step size and dwell time [20]. 8. The step-finding algorithm consists of three processes [20]: (i) finding steps, (ii) evaluating the quality of the step fits, and (iii) finding the step distribution. (i) The method of finding steps is not modified from the original [20]. Briefly, by fitting a single large step to the data, the first step is determined based on a calculation of the c2 statistics. Next, the subsequent steps are found by fitting new steps to the dwells of the previous fitting,

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in the same way as for the first step. (ii) The second process was customized for the cell tracking data as follows: the quality of fitting was evaluated by the variance   2in each Pof displacement dwell, which was defined as Q ¼ 1= iN þ1 σ 2i
σ 2 , where N is the number of steps, σ 2i is the variance of displacement in the i-th dwell, and hσ 2i is the mean of the variances in each b ¼ wQ ) was dwell. In addition, Q weighed by w ¼ 1/hσ 2i (Q b used for the third process. (iii) The highest Q value basically b, represents the best fitting. In practical terms, a value of Q which was not the highest value (usually the second highest value) toward the overfitting was often adopted as the best fitting, especially when the data showed a nonuniform fluctuation of the displacement. 9. High concentration [0.013% (w/v)] of Triton X-100 was used to observe the rotary movement of ghosts. Under this conditions, we detected gliding movements in 50% of the ghosts and rotary movements around the center like a tethered flagellated bacterial cell in a few percentage of ghosts [25–27]. This rotary movement occurred in both directions (56% in CW direction and 44% in CCW direction). We analyzed these movements and proposed a pivot model for this rotation, where the legs produce the thrust for the rotation and a flexible point, such as a membrane, is anchored to the surface (see the details in [28]). 10. The force generation process comprises both the chemical reaction of nucleotide and the binding process of leg proteins. You can control the latter by addition sialyllactose. 11. The binarization process should be applied by a threshold before the centroid position is measured by fitting. Various types of plug-ins are available in this process for Image J. References 1. Miyata M, Robinson RC, Uyeda TQP et al (2020) Tree of motility – a proposed history of motility systems in the tree of life. Genes Cells 25(1):6–21. https://doi.org/10.1111/ gtc.12737 2. Morio H, Kasai T, Miyata M (2016) Gliding direction of Mycoplasma mobile. J Bacteriol 198(2):283–290. https://doi.org/10.1128/ jb.00499-15 3. Jaffe JD, Stange-Thomann N, Smith C et al (2004) The complete genome and proteome of Mycoplasma mobile. Genome Res 14(8): 1447–1461. https://doi.org/10.1101/gr. 2674004 4. Uenoyama A, Miyata M (2005) Identification of a 123-kilodalton protein (Gli123) involved in machinery for gliding motility of Mycoplasma mobile. J Bacteriol 187(16):

5578–5584. https://doi.org/10.1128/jb. 187.16.5578-5584.2005 5. Adan-Kubo J, Uenoyama A, Arata T et al (2006) Morphology of isolated Gli349, a leg protein responsible for Mycoplasma mobile gliding via glass binding, revealed by rotary shadowing electron microscopy. J Bacteriol 188(8):2821–2828. https://doi.org/10. 1128/jb.188.8.2821-2828.2006 6. Kasai T, Nakane D, Ishida H et al (2013) Role of binding in Mycoplasma mobile and Mycoplasma pneumoniae gliding analyzed through inhibition by synthesized sialylated compounds. J Bacteriol 195(3):429–435. https:// doi.org/10.1128/jb.01141-12 7. Uenoyama A, Kusumoto A, Miyata M (2004) Identification of a 349-kilodalton protein (Gli349) responsible for cytadherence and

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glass binding during gliding of Mycoplasma mobile. J Bacteriol 186(5):1537–1545. h t t p s : // d o i . o r g / 1 0 . 1 1 2 8 / j b . 1 8 6 . 5 . 1537-1545.2004 8. Uenoyama A, Seto S, Nakane D et al (2009) Regions on Gli349 and Gli521 protein molecules directly involved in movements of Mycoplasma mobile gliding machinery, suggested by use of inhibitory antibodies and mutants. J Bacteriol 191(6):1982–1985. https://doi. org/10.1128/jb.01012-08 9. Kobayashi K, Kodera N, Kasai T et al (2021) Movements of Mycoplasma mobile gliding machinery detected by high-speed atomic force microscopy. mBio 12(3):e0004021. https://doi.org/10.1128/mBio.00040-21 10. Nonaka T, Adan-Kubo J, Miyata M (2010) Triskelion structure of the Gli521 protein, involved in the gliding mechanism of Mycoplasma mobile. J Bacteriol 192(3):636–642. https://doi.org/10.1128/jb.01143-09 11. Seto S, Uenoyama A, Miyata M (2005) Identification of a 521-kilodalton protein (Gli521) involved in force generation or force transmission for Mycoplasma mobile gliding. J Bacteriol 187(10):3502–3510. https://doi.org/10. 1128/jb.187.10.3502-3510.2005 12. Nakane D, Miyata M (2007) Cytoskeletal “jellyfish” structure of Mycoplasma mobile. Proc Natl Acad Sci U S A 104(49):19518–19523. https://doi.org/10.1073/pnas.0704280104 13. Nishikawa MS, Nakane D, Toyonaga T et al (2019) Refined mechanism of Mycoplasma mobile gliding based on structure, ATPase activity, and sialic acid binding of machinery. mBio 10(6). https://doi.org/10.1128/mBio. 02846-19 14. Toyonaga T, Kato T, Kawamoto A et al (2021) Chained structure of dimeric F1-like ATPase in Mycoplasma mobile gliding machinery. mBio 12(4):e0141421. https://doi.org/10.1128/ mBio.01414-21 15. Tulum I, Yabe M, Uenoyama A et al (2014) Localization of P42 and F1-ATPase α-subunit homolog of the gliding machinery in Mycoplasma mobile revealed by newly developed gene manipulation and fluorescent protein tagging. J Bacteriol 196(10):1815–1824. https:// doi.org/10.1128/jb.01418-13 16. Uenoyama A, Miyata M (2005) Gliding ghosts of Mycoplasma mobile. Proc Natl Acad Sci U S A 102(36):12754–12758. https://doi.org/ 10.1073/pnas.0506114102 17. Veigel C, Schmidt CF (2011) Moving into the cell: single-molecule studies of molecular motors in complex environments. Nat Rev Mol

Cell Biol 12(3):163–176. https://doi.org/10. 1038/nrm3062 18. Kinosita Y, Nakane D, Sugawa M et al (2014) Unitary step of gliding machinery in Mycoplasma mobile. Proc Natl Acad Sci U S A 111(23):8601–8606. https://doi.org/10. 1073/pnas.1310355111 19. Thompson RE, Larson DR, Webb WW (2002) Precise nanometer localization analysis for individual fluorescent probes. Biophys J 82(5): 2775–2783. https://doi.org/10.1016/ s0006-3495(02)75618-x 20. Kerssemakers JW, Munteanu EL, Laan L et al (2006) Assembly dynamics of microtubules at molecular resolution. Nature 442(7103): 7 0 9 – 7 1 2 . h t t p s : // d o i . o r g / 1 0 . 1 0 3 8 / nature04928 21. Kohori A, Chiwata R, Hossain MD et al (2011) Torque generation in F1-ATPase devoid of the entire amino-terminal helix of the rotor that fills half of the stator orifice. Biophys J 101(1):188–195. https://doi.org/10.1016/j. bpj.2011.05.008 22. Kitamura K, Tokunaga M, Iwane AH et al (1999) A single myosin head moves along an actin filament with regular steps of 5.3 nanometres. Nature 397(6715):129–134. https:// doi.org/10.1038/16403 23. Schnitzer MJ, Block SM (1997) Kinesin hydrolyses one ATP per 8-nm step. Nature 388(6640):386–390. https://doi.org/10. 1038/41111 24. Svoboda K, Schmidt CF, Schnapp BJ et al (1993) Direct observation of kinesin stepping by optical trapping interferometry. Nature 365(6448):721–727. https://doi.org/10. 1038/365721a0 25. Berg HC (1974) Dynamic properties of bacterial flagellar motors. Nature 249(452):77–79. https://doi.org/10.1038/249077a0 26. Larsen SH, Reader RW, Kort EN et al (1974) Change in direction of flagellar rotation is the basis of the chemotactic response in Escherichia coli. Nature 249(452):74–77. https://doi. org/10.1038/249074a0 27. Silverman M, Simon M (1974) Flagellar rotation and the mechanism of bacterial motility. Nature 249(452):73–74. https://doi.org/10. 1038/249073a0 28. Kinosita Y, Miyata M, Nishizaka T (2018) Linear motor driven-rotary motion of a membrane-permeabilized ghost in Mycoplasma mobile. Sci Rep 8(1):11513. https://doi.org/ 10.1038/s41598-018-29875-9

Chapter 28 Direct Measurement of Kinetic Force Generated by Mycoplasma Masaki Mizutani and Makoto Miyata Abstract Optical tweezers enable us to measure the force generated by bacterial motility and motor proteins. Here, we describe a method, using optical tweezers and related techniques, to measure the force generated during Mycoplasma gliding. An avidin-conjugated polystyrene bead trapped by a focused laser beam is bound to the surface-biotinylated Mycoplasma cell, which pulls the bead from the trap center of the laser. The force generated by Mycoplasma is calculated from a displacement measured and a spring constant of the laser trap. Key words Optical tweezers, Stall force, Avidin-biotin, Gliding motility, Bead, Pathogenic bacteria

1

Introduction Force is a crucial factor for understanding and modeling the motility of a cell. Optical tweezers are traditionally used to measure the kinetic force generated by bacterial motility and motor proteins [1– 7]. They were developed by Arthur Ashkin, who won the Nobel Prize in physics in 2018 [8]. Optical tweezers are a combination of an optical microscope and a laser. The laser beam focused by an objective lens can trap micro-sized molecules with a piconewtonorder force comparable to the force generated by bacterial motilities [3, 4, 9–11]. Here, we focused on the gliding motility of two bacterial species in the genus Mycoplasma. Mycoplasma are parasitic and occasionally commensal bacteria characterized by a small cell size, small genomes, and the absence of a peptidoglycan layer. Dozens of Mycoplasma species show gliding motility on solid surfaces, and this is thought to be essential for their virulence [12– 15]. Two types of gliding motility known in Mycoplasma are represented by M. mobile and M. pneumoniae [14–16], which have no homology in their component proteins. In this chapter, we describe detailed protocols to measure the force generated during the motility of these two Mycoplasma species.

Tohru Minamino et al. (eds.), Bacterial and Archaeal Motility, Methods in Molecular Biology, vol. 2646, https://doi.org/10.1007/978-1-0716-3060-0_28, © The Author(s), under exclusive license to Springer Science+Business Media, LLC, part of Springer Nature 2023

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Materials Optical Tweezers

The setup is briefly summarized in Fig. 1, and its important components are listed below: 1. Inverted microscope. 2. Nd:YAG laser (λ = 1064 nm, power > 1 W). 3. Plano convex lens with lens holder. 4. Mirror (placed in the microscope). 5. High numerical aperture objective lens (NA > 1.4). 6. Piezoelectric stage controlled by a stage controller. 7. High-speed camera (>2000 frames per second). 8. Neutral density (ND) filter with filter holder. 9. Stage heater and lens heater.

2.2

Bead

1. Carboxylated polystyrene bead (Φ = 1.0 μm). 2. Coupling Buffer (50 mM MES [pH 5.2], 0.05% Proclin 300) for protein coupling. 3. EDAC [1-Ethyl-3-(3-Dimethylaminopropyl)carbodiimide] for protein coupling. 4. Avidin from egg white. 5. Desktop ultrasonic cleaner. 6. Phosphate buffer saline (PBS): 75 mM sodium phosphate, pH 7.3, 68 mM NaCl.

Fig. 1 Optical tweezers system. The laser beam irradiated with the Nd:YAG laser oscillator is spread by a plano convex lens, inserted into a microscope, reflected by a mirror, and focused on the inside of tunnel chamber by the high NA objective lens

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2.3

Tunnel Chamber

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1. Large coverslip (40 × 22 mm). 2. Small coverslip (18 × 18 mm). 3. Double-sided tape (5 mm wide). 4. Ethanol (99.5%). 5. Potassium hydroxide (KOH). 6. Coverslip rack for wash. 7. PBS. 8. Horse serum heat-treated at 56 °C for 30 min to inactivate complements. 9. Bovine serum albumin (BSA). 10. Glucose.

2.4 Cells and Cultivation

1. Mycoplasma mobile 163K (ATCC43663). 2. Mycoplasma pneumoniae M129 (ATCC29342). 3. Tissue culture flask (25 cm2, 75 cm2).

2.5

Aluotto Medium

1. Mix 2.1 g of heart infusion broth, 0.56 g of yeast extract, double distilled water, and 10 M NaOH to a final volume of 89.8 mL and adjust the solution at pH 7.5. 2. Autoclave the solution for 15 min at 121 °C. 3. Add 10 mL of heat-inactivated horse serum, 100 μL of 2.5 mg/mL amphotericin B, and 100 μL of 50 mg/mL ampicillin sodium. 4. Store at 4 °C.

2.6

SP-4 Medium

1. Solution 1: Mix 0.35 g of Mycoplasma broth base (e.g., BD Biosciences, NJ, USA), 1 g of tryptone, 0.53 g of peptone, double distilled water, and 4 M KOH to a final volume of 60 mL and pH 7.7. 2. Autoclave the solution 1 for 15 min at 121 °C. 3. Solution 2: Mix 2.5 mL of 20% glucose, 5 mL of CMRL1066 (10×) w/o L-gutamine, 1.5 mL of 7.5% sodium bicarbonate, 0.5 mL of 200 mM L-gutamine, 3.5 mL of 25% fresh yeast extract solution, 10 mL of 2% TC yeastolate, 17 mL of heatinactivated fetal bovine serum (Gibco™; Thermo Fisher Scientific, MA, USA), 0.25 mL of 400,000 U/mL penicillin G, and 0.15 mL of 1% phenol red. 4. Pass solution 2 through a 0.22 μm pore size filter for sterilization—a process known as filter sterilization. 5. Mix 60 mL of solution 1 and 40 mL of solution 2. 6. Store at 4 °C.

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2.7 Surface Modification and Preparation of Cells

1. PBS. 2. Sulfo-NHS-LC-LC-biotin (EZ-Link™ Sulfo-NHS-LC-LCbiotin; Thermo Fisher Scientific). 3. Glucose. 4. Polyvinylidene difluoride (PVDF) membrane filter (pore size: 0.45 μm) and syringe. 5. Horse serum (not heat-inactivated).

3

Methods

3.1 Cultivation of Mycoplasma mobile

1. Inoculate 1 mL of a frozen stock into 10 mL of Aluotto medium in 25 cm2 of tissue culture flask. 2. Incubate the culture statically at 25 °C to an optical density of 0.06–0.08 at 600 nm.

3.2 Cultivation of Mycoplasma pneumoniae

3.3 Biotin Conjugation to Mycoplasma mobile Cell Surface

1. Inoculate 1 mL of a frozen stock into 30 mL of SP-4 medium in 75 cm2 of tissue culture flask. 2. Incubate the culture statically at 37 °C until the color of culture changes to orange (see Note 1). 1. Collect the cells via centrifugation at 12,000 × g for 4 min at 25 °C. 2. Wash the cells with 3 mL of PBS. 3. Suspend the cells in 450 μL of PBS containing 0.5 mM of Sulfo-NHS-LC-LC-biotin. 4. Incubate the cells for 15 min at 25 °C. 5. Collect the cells via centrifugation at 12,000 × g for 4 min at 25 °C. 6. Wash the cells with 1.5 mL of PBS twice. 7. Suspend the cells in 450 μL of PBS containing 20 mM glucose. 8. Keep the cell suspension at 25 °C before use.

3.4 Biotin Conjugation to Mycoplasma pneumoniae Cell Surface

1. Gently rinse the cultured cells with 4 mL of PBS twice in the culture flask (see Note 2). 2. Add 1 mL of PBS mixed with 0.5 mM of Sulfo-NHS-LC-LCbiotin. 3. Collect the cells by scraping them from the bottom of flask. 4. Pass the cell suspension through a filter with pores sized 0.45 μm. 5. Incubate for 15 min at 25 °C. 6. Collect the cells via centrifugation at 12,000 × g for 10 min at 25 °C.

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7. Wash the cells with 1.5 mL of PBS twice. 8. Suspend the cells in 500 μL of PBS containing 10% horse serum (non-heat-inactivated) and 20 mM glucose (see Note 3). 9. Pass the cell suspension through a filter with pores sized 0.45 μm. 10. Keep the cell suspension at 37 °C before use. 3.5 Avidin Conjugation to Polystyrene Beads

1. Collect polystyrene beads from the 50–100 μL of solution via centrifugation at 2000 × g for 6 min at 25 °C. 2. Suspend the beads in 400 μL of coupling buffer. 3. Centrifuge the suspension again at 2000 × g for 6 min at 25 °C. 4. Suspend the beads in 160 μL of coupling buffer. 5. Add 20 μL of 1 M EDAC to the suspension. 6. Incubate the suspension for 5 min at 25 °C. 7. Add 10 μL of 0.1 mM avidin into the suspension. 8. Incubate, with gentle turnover, for 3–5 h at 25 °C. 9. Wash the beads with PBS at least five times. 10. Store at 4 °C.

3.6 Construction and Surface Coating of Tunnel Chamber

1. Set large coverslips on the coverslip rack. 2. Prepare a fresh saturated potassium hydroxide ethanol solution by adding 80 g of potassium hydroxide to 350 ml of 99.5% ethanol in a 500 ml beaker. 3. Soak the coverslips in the saturated potassium hydroxide ethanol solution for 10 min at 25 °C. 4. Rinse the coverslips with distilled water ten times. 5. Dry the coverslips. 6. Stick two pieces of double-sided tape on the cleaned large coverslip (Fig. 2). 7. Stick a small coverslip on the tape. 8. Place the heat-inactivated horse serum in the tunnel chamber. 9. Incubate for 60 min at 25 °C (see Notes 4 and 5). 10. Replace with 20 μL of PBS containing 10 mg/mL BSA. 11. Incubate for 60 min at 25 °C (see Notes 4 and 6). 12. Replace with 20 μL of PBS containing 20 mM glucose. 13. Keep at 25 °C before use (see Note 4).

3.7 Measurements of Spring Constant of Optical Tweezers

1. Dilute the carboxylated polystyrene bead solution using PBS to avoid capturing multiple beads with the optical tweezers (see Note 7). 2. Insert the bead solution into a tunnel chamber.

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Fig. 2 Tunnel chamber construction. Two coverslips are combined with double sided tapes. Detailed methods are given in Subheading 3.6

3. Set the tunnel chamber on the microscope stage equipped with optical tweezers (see Note 8). 4. Launch the laser and reduce it to 1–4 mW by ND filter (see Note 9). 5. Trap a bead a few hundred nanometers above the lower coverslip and record its Brownian motion for 5 s at 2000 frames per second using the high-speed camera. 3.8 Data Analysis for Spring Constant Measurement

1. Trace the center of bead and create individual distribution histograms of the X and Y positions (Fig. 3). 2. Fit the histograms using the following function. 
 F ðx Þ = A  exp - ðx - x0Þ2 = 2 σ 2 where A is the peak of histogram, x0 is the center of histogram, and σ is the standard deviation of bead existence distribution. 3. Calculate the spring constant K using the following equation. K = kB  T =σ 2 =W where kB is the Boltzmann constant, T is the absolute temperature, and W is the laser power.

3.9 Measuring Force Using Optical Tweezers

1. Insert the biotinylated Mycoplasma cells into a precoated tunnel chamber. 2. Incubate Mycoplasma mobile at 25 °C and Mycoplasma pneumoniae at 37 °C for 15 min. 3. Remove floating cells by adding 20 μL of PBS containing 20 mM glucose.

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Fig. 3 Spring constant calculation. (a) Optical micrograph of polystyrene bead. Bar, 0.5 μm. (b) Trace of Brownian motion of the trapped bead. (c, d) Distribution histograms of the bead in Brownian motion. The center of distribution is already calibrated as 0

4. Dilute avidin-conjugated beads solution using PBS containing glucose (final concentration of 20 mM), and sonicate it (see Notes 7 and 10). 5. Insert 20 μL of avidin-conjugated beads into the tunnel chamber. 6. Enclose the entrance and exit of tunnel chamber with nail polish (see Note 11). 7. Set the tunnel chamber on the microscope stage equipped optical tweezers. 8. Incubate Mycoplasma mobile and Mycoplasma pneumoniae for a few min on the microscope stage at 25 °C and 37 °C, respectively. 9. Launch the laser (see Note 12). 10. Trap the bead at the near position of the lower coverslip and start to record with the camera. 11. Bind the bead to the back side of gliding cell body by moving the microscope stage and stop the stage movement just after binding (Fig. 4). 12. Continue to record until the gliding stalls.

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Fig. 4 Force measurements. (a) Experimental design. The avidin-conjugated polystyrene bead trapped by a focused laser beam is bound to the surface-biotinylated Mycoplasma cell. The cell glides on the serum-coated coverslip in the direction of the arrow. (b) Optical micrograph of a cell trapped by the bead (large black ring with white center). The trap center is marked by a red cross. Bar, 0.5 μm 3.10 Data Analysis for Force Measurement

1. Trace the center of bead and construct the coordinate table with its X and Y positions, individually, over time. 2. Calibrate the first position as 0. 3. Calculate the force using the following equation. h i1=2 F = ðX  Kx  W Þ2 þ ðY  Ky  W Þ2 where F is the force, Kx is the spring constant of X axis, Ky is the spring constant of Y axis, and W is the laser power.

4

Notes 1. The optical density of the culture is not useful for checking the growth of Mycoplasma pneumoniae because the cells grow on the bottom of the culture flask; phenol red, a pH indicator, is more useful [17]. 2. Some M. pneumoniae cells bind to the bottom of the culture flask. Strong shaking should be avoided to ensure cell attachment. 3. Horse serum containing active complements activates the gliding motility of M. pneumoniae [18, 19]. 4. The tunnel chamber was placed in a humid area to prevent the solution from evaporating. 5. Horse serum contains glycoproteins and glycolipids modified by sialic acids, which are the binding targets of Mycoplasma cells; it thus acts as a scaffold for gliding. 6. BSA blocks nonspecific binding between the cell body and glass surface.

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7. To measure the force, optical tweezers must capture a single bead. The dilution rate depends on the original concentration of the bead solution. 8. The temperature of the microscope stage should be maintained using stage and lens heaters at 25 °C to measure the force of M. mobile and at 37 °C for M. pneumoniae. 9. The laser emissions are sometimes unstable at low power. 10. Immediately before insertion, the bead solution was sonicated because beads easily bind to each other. 11. The enclosure of the tunnel chamber prevents the solution from evaporating and any subsequent water flow, which is harmful for measurements [20]. 12. To measure the force accurately, bead displacements should be limited to 200–250 nm from the trap center [9]. The laser power should be regulated to keep a bead in the range (e.g., 600 mW for M. mobile, 250 mW for M. pneumoniae).

Acknowledgments We thank Takayuki Nishizaka at Gakushuin University for designing and constructing optical tweezers. This work was supported by Grants-in-Aid for Scientific Research (A) (MEXT KAKENHI, Grant Number JP17H01544) and by JST CREST (Grant Number JPMJCR19S5, Japan). References 1. Nishizaka T, Miyata H, Yoshikawa H et al (1995) Unbinding force of a single motor molecule of muscle measured using optical tweezers. Nature 377:251–254 2. Kojima H, Muto E, Higuchi H et al (1997) Mechanics of single kinesin molecules measured by optical trapping nanometry. Biophys J 73(4):2012–2022. https://doi.org/10. 1016/S0006-3495(97)78231-6 3. Merz AJ, So M, Sheetz MP (2000) Pilus retraction powers bacterial twitching motility. Nature 407(6800):98–102. https://doi.org/ 10.1038/35024105 4. Miyata M, Ryu WS, Berg HC (2002) Force and velocity of Mycoplasma mobile gliding. J Bacteriol 184:1827–1831. https://doi.org/ 10.1128/JB.184.7.1827-1831.2002 5. Clemen AE, Vilfan M, Jaud J et al (2005) Force-dependent stepping kinetics of myosin-V. Biophys J 88(6):4402–4410. https://doi.org/10.1529/biophysj.104. 053504

6. Takagi Y, Homsher EE, Goldman YE et al (2006) Force generation in single conventional actomyosin complexes under high dynamic load. Biophys J 90(4):1295–1307. https:// doi.org/10.1529/biophysj.105.068429 7. Gennerich A, Carter AP, Reck-Peterson SL et al (2007) Force-induced bidirectional stepping of cytoplasmic dynein. Cell 131(5): 952–965. https://doi.org/10.1016/j.cell. 2007.10.016 8. Ashkin A, Dziedzic JM, Bjorkholm JE et al (1986) Observation of a single-beam gradient force optical trap for dielectric particles. Opt Lett 11(5):288 9. Tanaka A, Nakane D, Mizutani M et al (2016) Directed binding of gliding bacterium, Mycoplasma mobile, shown by detachment force and bond lifetime. MBio 7(3):e00455–e00416. https://doi.org/10.1128/mBio.00455-16 10. Mizutani M, Tulum I, Kinosita Y et al (2018) Detailed analyses of stall force generation in Mycoplasma mobile gliding. Biophys J 114(6):

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1411–1419. https://doi.org/10.1016/j.bpj. 2018.01.029 11. Mizutani M, Sasajima Y, Miyata M (2021) Force and stepwise movements of gliding motility in human pathogenic bacterium Mycoplasma pneumoniae. Front Microbiol 12: 747905. https://doi.org/10.3389/fmicb. 2021.747905 12. Hatchel JM, Balish MF (2008) Attachment organelle ultrastructure correlates with phylogeny, not gliding motility properties, in Mycoplasma pneumoniae relatives. Microbiology 154(Pt 1):286–295. https://doi.org/10. 1099/mic.0.2007/012765-0 13. Prince OA, Krunkosky TM, Krause DC (2014) In vitro spatial and temporal analysis of Mycoplasma pneumoniae colonization of human airway epithelium. Infect Immun 82(2): 579–586. https://doi.org/10.1128/IAI. 01036-13 14. Miyata M, Hamaguchi T (2016) Prospects for the gliding mechanism of Mycoplasma mobile. Curr Opin Microbiol 29:15–21. https://doi. org/10.1016/j.mib.2015.08.010 15. Miyata M, Hamaguchi T (2016) Integrated information and prospects for gliding mechanism of the pathogenic bacterium Mycoplasma

pneumoniae. Front Microbiol 7:960. https:// doi.org/10.3389/fmicb.2016.00960 16. Miyata M, Robinson RC, Uyeda TQP et al (2020) Tree of motility – a proposed history of motility systems in the tree of life. Genes Cells 25(1):6–21. https://doi.org/10.1111/ gtc.12737 17. Terahara N, Tulum I, Miyata M (2017) Transformation of crustacean pathogenic bacterium Spiroplasma eriocheiris and expression of yellow fluorescent protein. Biochem Biophys Res Commun 487(3):488–493. https://doi.org/ 10.1016/j.bbrc.2017.03.144 18. Nakane D, Miyata M (2009) Cytoskeletal asymmetrical dumbbell structure of a gliding mycoplasma, Mycoplasma gallisepticum, revealed by negative-staining electron microscopy. J Bacteriol 191(10):3256–3264. https:// doi.org/10.1128/JB.01823-08 19. Mizutani M, Miyata M (2019) Behaviors and energy source of Mycoplasma gallisepticum gliding. J Bacteriol 201(19):e00397–e00319. https://doi.org/10.1128/JB.00397-19 20. Mizutani M, Miyata M (2017) Force measurement on Mycoplasma mobile gliding using optical tweezers. Bio Protoc 7:e2127. https://doi. org/10.21769/BioProtoc.2127

Chapter 29 Genetic Manipulation of Mycoplasma pneumoniae Tsuyoshi Kenri Abstract Mycoplasma pneumoniae is a small cell wall-lacking bacterium that is a common cause of bronchitis and pneumonia in humans. In addition to its clinical importance, M. pneumoniae has recently been considered a promising model organism for synthetic biology because of its small genome size and unique cell structure. At one cell pole, M. pneumoniae forms the attachment organelle that is responsible for adherence to host cells and gliding motility. The attachment organelle is a membrane protrusion and is composed of number of molecules, including adhesin and cytoskeletal proteins. Genetic manipulation techniques are key research approaches for understanding the structure and the function of this unique molecular machinery. In this chapter, standard genetic engineering methods for this species using the Tn4001 transposon vector are described. Key words Mycoplasma pneumoniae, Transposon, Tn4001, Gateway cloning, Transformation, Fluorescent protein, GFP, mCherry

1

Introduction Transformation of Mycoplasma pneumoniae by the Staphylococcus aureus transposon Tn4001 is a classical and standard genetic manipulation system for this bacterium. Using Tn4001 vectors, exogenous genes, such as fluorescent protein tags or reporter enzyme genes, can be efficiently introduced into M. pneumoniae cells [1, 2]. Unlike transformation using plasmids, the Tn4001 vector is randomly inserted at various locations in the genome; therefore, it is also a useful method for creating gene disruption mutants [3– 5]. Transformation of Mycoplasma species by using Tn4001 was first reported by Mahairas et al. [1], and this method was applied to M. pneumoniae by Hedreyda et al. [6]. Since these reports, many Tn4001 vectors have been designed and used for various applications [2, 7, 8]. In recent years, mini-Tn4001 vectors composed of a minimal number of elements (transposase genes, recombination sites, and selection markers) have also been developed [9, 10]. However, in experiments using Tn4001 vectors, the construction of

Tohru Minamino et al. (eds.), Bacterial and Archaeal Motility, Methods in Molecular Biology, vol. 2646, https://doi.org/10.1007/978-1-0716-3060-0_29, © The Author(s), under exclusive license to Springer Science+Business Media, LLC, part of Springer Nature 2023

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relatively large plasmids is required and is often time-consuming. The Gateway cloning system [11] is very useful for these experiments, enabling precise assembly of large Tn4001 plasmids for transformation experiments with M. pneumoniae. This chapter describes the construction of a standard Tn4001 plasmid that carries a large fusion gene for expression in M. pneumoniae using Gateway cloning, the method for transformation of M. pneumoniae by the Tn4001 plasmid via electroporation, selection procedure of a single transformant cell, and analysis of the Tn4001 insertion site on the transformant chromosome. This information will be useful for further molecular biology investigations and understanding of this unique bacterial pathogen [12–14].

2

Materials

2.1 Bacterial Strains and Plasmids

1. Mycoplasma pneumoniae M129 (ATCC 29342™). 2. Escherichia coli DB 3.1: F
, gyrA462, endA1, Δ(sr1-recA), mcrB, mrr, hsdS20, glnV44 (¼supE44), ara14, galK2, lacY1, proA2, rpsL20, xyl5, λ
, leuB6, mtl1. 3. Escherichia coli DH5α: F
, Φ80dlacZΔM15, Δ(lacZYAargF), U169, deoR, recA1, endA1, hsdR17(rK
, mK+), phoA, supE44, λ
, thi1, gyrA96, relA1. 4. The pKM310-ST (pKM310-standard) plasmid (GenBank accession No. LC671235) [15]. 5. The pTK170-D plasmid (GenBank accession No. LC671237) [15, 16]. 6. The pKM170-ST (pKM170-standard) plasmid (GenBank accession No. LC671236) [15].

2.2

Media

1. PPLO broth: 2.1% (w/v) Difco™ PPLO broth without CV, 0.25% (w/v) glucose, 0.002% (w/v) phenol red, 10% (v/v) horse serum, 0.25% (w/v) Bacto™ yeast extract, 1.25% (w/v) fresh yeast extract, and 50 mg/mL ampicillin. Store at 4  C (see Note 1). 2. PPLO agar plate: 3.5% (w/v) Difco™ PPLO agar, 0.25% (w/v) glucose, 10% (v/v) horse serum, 0.25% (w/v) Bacto™ yeast extract, 1.25% (w/v) fresh yeast extract, and 50 mg/mL ampicillin. Store at 4  C (see Note 1). 3. LB broth: 1% (w/v) Bacto™ Tryptone, 0.5% (w/v) Bacto™ yeast extract, 0.5% (w/v) NaCl, and 0.1% (w/v) glucose. 4. LB agar plates: 1% (w/v) Bacto™ Tryptone, 0.5% (w/v) Bacto™ yeast extract, 0.5% (w/v) NaCl, 0.1% (w/v) glucose, and 1.5% (w/v) agar. Stored at 4  C. 5. 50 mg/mL ampicillin: 1000 stock solution. Stored at
20  C. 6. 50 mg/mL kanamycin: 1000 stock solution. Stored at
20  C.

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7. 15 mg/mL chloramphenicol: 1000 stock solution (dissolved in ethanol). Store at
20  C. 8. 18 mg/mL gentamicin (potency): 1000 stock solution. Stored at
20  C. 9. Phosphate buffered saline (PBS): 137 mM NaCl, 3 mM KCl, 8 mM Na2HPO4, 1 mM KH2PO4, pH 7.4. 10. 75 cm2 cell culture flask. 11. 25 cm2 cell culture flask. 12. Cell scraper: TPP 99003. 13. 2.5 mL sterile syringe and 25-gauge needle. 14. 0.45 μm membrane filter. 2.3 Enzymes and Kits

1. LR clonase™ II Mix. 2. PrimeSTAR™ Max DNA polymerase. 3. QIAprep Spin Miniprep Kit. 4. QIAamp DNA Mini Kit. 5. DNA Ligation Kit. 6. HindIII. 7. 2 μg/μL proteinase K solution. 8. TE buffer: 10 mM Tris–HCl, 1 mM EDTA, pH 8.0.

2.4

Electroporation

1. Electroporation system (e.g., Gene Pulser Xcell, Bio-Rad). 2. Electroporation cuvettes, 0.2 cm gap (e.g., Gene Pulser Electroporation Cuvettes Bio-Rad). 3. Electroporation (EP) buffer: 8 mM HEPES, 272 mM sucrose, pH 7.4. Stored at 4  C. 4. Centrifuge tubes: 50 mL, 15 mL, 1.5 mL.

2.5 Oligo DNA Primers

3

1. 4001R: 50 -AAACATTGTACCGTAAAAGG-30 . 2. 4001IV: 50 -TTAACTTAGCGCGTGAGGCT-30 .

Methods

3.1 Construction of Tn4001 Vector Plasmid Using Gateway Cloning (See Note 2)

1. Inoculate E. coli DH5α harboring pKM310-ST plasmid (entry clone) (Fig. 1) into 5 mL of LB broth containing 50 μg/mL kanamycin. Inoculate E. coli DB3.1 harboring pTK170-D plasmid (destination vector) into 5 mL of LB broth containing 50 μg/mL ampicillin and 15 μg/mL chloramphenicol. Incubate at 37  C overnight with shaking. 2. Prepare pKM310-ST and pTK170-D plasmids using QIAprep Spin Miniprep Kit or similar products.

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Fig. 1 Scheme of construction of the Tn4001 vector plasmid using the Gateway cloning method. The pKM310ST plasmid (entry clone) carries the mcherry-hmw2(ΔP28) fusion gene between the attR1 and attL1 recombination sites [15]. The pTK170-D (destination vector) carries the tuf promoter of Mycoplasma pneumoniae (Ptuf) and eyfp gene in the SmaI site of Tn4001mod (pISM2062.2) [7, 16]. The CmR marker, ccd gene, and attR recombination sites were inserted between Ptuf and eyfp. Recombination by LR clonase generates the pKM170-ST plasmid that carries the mcherry-hmw2(ΔP28)-eyfp fusion gene under the control of Ptuf [15, 16]. When the Tn4001 vector plasmid enters M. pneumoniae cells, transposition of Tn4001 sequence (between IR-L and IR-R) into random sites of M. pneumoniae chromosome occurs and confers gentamicin resistance to M. pneumoniae

3. Mix 100 ng of pKM310-ST and 150 ng of pTK170-D in 8 μL of TE buffer. Add 2 μL of LR Clonase™ II Mix and vortex briefly. Incubate the mixture at 25  C for 1 h. 4. Add 1 μL of the proteinase K solution and vortex briefly. Incubate the mixture at 37  C for 10 min.

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5. Add 1–2 μL of LR reaction mixture into 100 μL of DH5α competent cell. Incubate the mixture tube on ice for 30 min. Heat-shock cells by placing the tube at 42  C for 30 s. Add 900 μL of LB broth and incubate at 37  C for 1 h. Plate 50–100 μL of transformation mixture onto selective LB plates containing 50 μg/mL of ampicillin. 6. Select several transformant colonies and inoculate into 3 mL of LB broth containing 50 μg/mL of ampicillin. Incubate at 37  C overnight with shaking. 7. Prepare the pKM170-ST plasmid using QIAprep Spin Miniprep Kit or similar products. Check the plasmid size (Fig. 1) using 0.8% (w/v) agarose gel electrophoresis. If needed, analyze the plasmid using PCR or other sequencing methods. 8. Prepare the pKM170-ST plasmid on large scale. 3.2 Transformation of M. pneumoniae

1. Inoculate 100 μL of frozen stock of M. pneumoniae M129 strain (or other strain) in 5 mL of PPLO broth and incubate the culture at 37  C for several days until the PPLO broth color turns orange (mid-log phase). 2. Inoculate 1–2 mL of the preculture into 100 mL of fresh PPLO broth in a 75 cm2 tissue culture flask. Place the flasks on their sides in the incubator to rest. Incubate at 37  C for 2–5 days until the PPLO broth color turns orange (mid-log phase). 3. Before the preparation of M. pneumoniae competent cells, place the EP buffer, fresh PPLO broth, centrifuge tubes (15 mL and 1.5 mL), and electroporation cuvettes on ice for approximately 20 min. 4. Discard the PPLO culture broth and add 20 mL of ice-cold EP buffer to the culture flask. Wash the inside of the flask by gentle shaking and discard the EP buffer. Repeat this step once (see Note 3). 5. Add 5 mL of ice-cold EP buffer to the flask and harvest M. pneumoniae cells from the sidewall of the flask (bottom side during culture) using a cell scraper. 6. Collect the cell suspension in a 15 mL centrifuge tube and place it on ice until ready to use (see Note 4). 7. Set the electroporation machine parameters as follows: 2.5 kV, 100 Ω resistance, and 25 μF. 8. Add 0.5–5 μg of pKM170-ST plasmid DNA (approximately 1–10 μL of volume) to 100 μL of cell suspension in a 1.5 mL centrifuge tube. Gently vortex it and place it on ice for 10 min. For the negative control, add TE buffer (same volume as that of the DNA solution) to the cell suspension under the same conditions (see Note 5).

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9. Transfer the cells to a cold electroporation cuvette and tap the suspension to the bottom. 10. Place the cuvette in the electroporation chamber, pulse once (2.5 kV, 100 Ω resistance, 25 μF) by pushing the pulse button of the machine. 11. Remove the cuvette from the chamber and immediately add 1 mL of ice-cold PPLO broth. Mix with gentle shaking and incubate at 37  C for 2 h (see Note 6). 12. Plate 100–150 μL of the cell mixture on a PPLO agar plate containing 18 μg/mL gentamicin (see Note 7). Inoculate the remaining mixture into 10 mL of PPLO broth containing 18 μg/mL gentamicin. Incubate the PPLO plate and broth at 37  C for 1–2 weeks (see Note 8). 13. Observe the colonies on the PPLO plate using a stereomicroscope with 10–100 magnification (see Note 9). 14. Check the transformant growth in the PPLO broth by observing the color change of the broth (orange-yellow color) (see Note 10). Transfer the PPLO broth of transformant into 1–2 mL plastic vials and store at
80  C (see Note 11) (Fig. 2).

Fig. 2 Phase-contrast and fluorescence microscopy of the transformant cells that produce the mCherryHMW2-EYFP fusion protein. (a) Phase contrast, mCherry, EYFP, and merged images of transformant cells. Bar ¼ 2 μm (b) Enlarged merged image of the cells and schematic illustration of the attachment organelle. The attachment organelle is a membrane protrusion at one cell pole of M. pneumoniae [13]. Adhesin protein complexes consisting of the P1, P40, and P90 proteins are present at the organelle surface [18]. The organelle is supported by an internal cytoskeleton-like structure (core). The HMW2 proteins are major components of the core and are aligned in parallel along the longer axis of the core with the N-terminus at the front as demonstrated by fluorescence signals from mCherry-HMW2-EYFP fusion protein [15, 16]

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Fig. 3 Mycoplasma pneumoniae single-colony cloning method. (a) M. pneumoniae single-colony selection on a PPLO agar plate using a 200 μL yellow pipette tip. The colony and agar were punched by the tip. The arrow indicates the colony. (b) Punched agar and a single colony in PPLO broth. The arrow indicates the colony. (c) Magnified side view of an M. pneumoniae colony. M. pneumoniae cells enter the agar during growth and form colonies 3.3 Analysis of Tn4001 Insertion Site (Inverse PCR Method)

1. Pick a single transformant colony from a PPLO agar plate using a sterile disposable pipette tip under a stereomicroscope (Fig. 3) and inoculate into 2–3 mL of fresh PPLO broth containing 18 μg/mL gentamicin. Incubate at 37  C until the mid-log phase, that is, the PPLO broth color turns orange (usually 5–14 days). 2. Using a 2.5 mL sterile syringe, pass the PPLO culture through a 25-gauge needle several times, then pass it through a 0.45 μm membrane filter, and serially dilute it (tenfold) to 10
6. 3. Place 5–10 μL of each dilution on the PPLO agar plate containing 18 μg/mL gentamicin and incubate at 37  C for 5–14 days. 4. Repeat the steps from procedure 1 once or twice when colonies are formed on the plate. 5. Pick a single transformant colony on a PPLO agar plate, inoculate the colony into 2–3 mL of fresh PPLO broth containing 18 μg/mL gentamicin, and incubate at 37  C until the mid-log phase. 6. Inoculate 0.5–1 mL of the preculture into 10 mL of fresh PPLO broth in a 25 cm2 tissue culture flask. Store the remaining preculture at -80  C. Place the culture flasks on their sides in the incubator to rest. Incubate at 37  C for 2–5 days until the mid-log phase is attained.

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7. Discard PPLO broth and add 1 mL of PBS to the flask. Harvest M. pneumoniae cells from the sidewall of the flask (bottom side during culture) using a cell scraper (see Note 12) and place them into a 1.5 mL centrifugation tube. 8. Centrifuge at 20,000  g for 10 min at 4  C to obtain the cell pellet. Discard the PBS and resuspend the cells in 0.5 mL of PBS. 9. Extract the genomic DNA from 200 μL of the cell suspension using the QIAamp DNA Mini Kit or a similar product. 10. Measure the absorbance at 260 nm and estimate the DNA concentration. Check the quality of the genomic DNA using 0.8% (w/v) agarose gel electrophoresis or other methods. 11. Digest completely 1 μg of purified genomic DNA using restriction endonuclease HindIII. 12. Self-ligate 0.5 μg of HindIII -digested DNA using a DNA ligation kit or T4 DNA ligase (in total 20 μL reaction mixture). After ligation, clean the DNA using ethanol precipitation. 13. Amplify the DNA via PCR using 50 ng of self-ligated DNA as a template and the primer set 4001R: 50 -AAACATTGTACCG TAAAAGG -30 and 4001IV: 50 - TTAACTTAGCGCGT GAGGCT-30 (see Note 13). 14. Check the PCR product using 0.8% (w/v) agarose gel electrophoresis. If there are multiple PCR products, try another PCR condition, such as purification of the major products using gel extraction or subcloning the products into a plasmid vector. 15. Sequence the PCR products using the 4001R primer. 16. Using the obtained sequences, find and analyze the IR-R side of Tn4001 and the insertion site (Fig. 4) (see Note 14). 17. If needed, confirm the Tn4001 insertion site using another specific PCR primer set or Southern blotting experiment of genomic DNA of the transformant.

4

Notes 1. Add 25% fresh yeast extract (an aqueous extract of baker’s yeast) to the media at a final concentration of 1.25% (one twentieth volume). A 25% fresh yeast extract is prepared as reported previously by Hayflick [17]. It is also commercially available from several companies (e.g., Oriental Yeast, Japan). 2. This is the procedure for constructing the pKM170-ST plasmid [15] and is an example of Tn4001 vector construction using Gateway cloning (Fig. 1). The pKM170-ST plasmid carried the mcherry-hmw2(ΔP28)-eyfp fusion gene in Tn4001mod [7] for

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Fig. 4 The Tn4001 insertion site on M. pneumoniae transformant chromosome. Sequence of the right end (30 end) of Tn4001 is shown. The inverted repeat of Tn4001 (IR-R) and the 4001R primer binding site are indicated. The inverse PCR products contain this region (boundary between Tn4001 and M. pneumoniae chromosome DNA sequences). The insertion site is specified by sequencing of the PCR product using the 4001R primer. The information of the chromosome sequence adjacent to the 30 end of Tn4001 is utilized for 50 end analysis (see Note 14)

expression in M. pneumoniae cells. The pTK170-D plasmid [15, 16] is a destination vector for Gateway cloning based on the standard Tn4001mod vector, pISM2062.2 [7]. The conversion of the Tn4001mod vector into Gateway destination vectors enables accurate and efficient incorporation of large gene fragments into the Tn4001mod using entry clone cassettes such as pKM310-ST [15]. 3. In the case of nonadherent M. pneumoniae cells (hemadsorption negative mutants), harvest the cells using centrifugation at 20,000  g for 20 min at 4  C. Resuspend the pellet in 50 mL of EP buffer and wash the cells. Centrifuge at 20,000  g for 20 min at 4  C and resuspend cells in 5 mL of EP buffer. 4. At this point, the CFU of the cell suspension is usually around 108 to 109/mL, depending on the culture conditions and strain. 5. Add sterile 80% (v/v) glycerol to the remaining competent cell to a final glycerol concentration of 10% (w/v), and store at
80  C for the next experiment. The competent cells can be used for several months. 6. Alternatively, transfer the cells from the cuvette to 1 mL of cold PPLO in a 1.5 mL centrifuge tube on ice using a thin pipette and then incubate the tube at 37  C for 2 h. 7. To calculate the transformation efficiency (gentamicin-resistant CFU/total viable CFU), plate the diluted cell mixture (10
1 to 10
6) on the PPLO agar plates not containing selection drugs. 8. For comparison of medium color, incubate PPLO broth only (at least one tube) under the same conditions, as the color of PPLO broth slightly changes during the culture period without the growth of M. pneumoniae.

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9. Note that no colony formation occurred on the negative control plate (without plasmid DNA). Count the colonies and calculate the transformation efficiency (CFU/μg DNA or gentamicin-resistant CFU/total viable CFU). 10. Note that no color change occurred in the negative control tube (without plasmid DNA) compared to that of PPLO broth only. 11. The transformant cells selected in the PPLO broth are a mixture, and the chromosomal location of the Tn4001 insertion site varies between cells. Selecting a single colony on an agar plate is needed to obtain a clonal transformant strain. Transformant cells in this experiment example exhibit mCherry and EYFP fluorescence at the attachment organelle, which is present at one cell pole of this bacterium (Fig. 2) [14]. 12. For nonadherent centrifugation.

mutants,

harvest

the

cells

using

13. For PrimeSTAR™ Max DNA polymerase, the reaction conditions are 98  C 10 s; 30 cycles of 98  C 10 s; 58  C 10 s; 72  C 30 s; 72  C 3 min. 14. In addition, design the specific PCR primer sets of the insertion site using the information of the chromosome sequence adjacent to the IR-R of Tn4001 (Fig. 4) and the Tn4001 vector sequence (GenBank: LC671236). Using the primer sets and the genomic DNA of the transformant, amplify the DNA region containing the IR-L side of Tn4001, sequence the DNA, and confirm the insertion site.

Acknowledgments This work was supported by Grants-in-Aid for Scientific Research (JP25117530 and JP15H01337) from the Ministry of Education, Culture, Sports, Science, and Technology of Japan and grants from the Japan Agency for Medical Research and Development (21jk0210004j0101 and 20jk0210004j0101). References 1. Mahairas GG, Minion FC (1989) Random insertion of the gentamicin resistance transposon Tn4001 in Mycoplasma pulmonis. Plasmid 21(1):43–47. https://doi.org/10.1016/ 0147-619x(89)90085-1 2. Kenri T, Seto S, Horino A et al (2004) Use of fluorescent-protein tagging to determine the subcellular localization of Mycoplasma pneumoniae proteins encoded by the cytadherence regulatory locus. J Bacteriol 186(20):6944–6955.

h t t p s : // d o i . o r g / 1 0 . 1 1 2 8 / J B . 1 8 6 . 2 0 . 6944-6955.2004 3. Krause DC, Proft T, Hedreyda CT et al (1997) Transposon mutagenesis reinforces the correlation between Mycoplasma pneumoniae cytoskeletal protein HMW2 and cytadherence. J Bacteriol 179(8):2668–2677. https://doi. org/10.1128/jb.179.8.2668-2677.1997 4. Reddy SP, Rasmussen WG, Baseman JB (1996) Isolation and characterization of transposon

Transformation of M. pneumoniae by Tn4001 Tn4001-generated, cytadherence-deficient transformants of Mycoplasma pneumoniae and Mycoplasma genitalium. FEMS Immunol Med Microbiol 15(4):199–211. https://doi.org/ 10.1111/j.1574-695X.1996.tb00086.x 5. Hutchison CA, Peterson SN, Gill SR et al (1999) Global transposon mutagenesis and a minimal Mycoplasma genome. Science (New York, NY) 286(5447):2165–2169. https://doi.org/10.1126/science.286.5447. 2165 6. Hedreyda CT, Lee KK, Krause DC (1993) Transformation of Mycoplasma pneumoniae with Tn4001 by electroporation. Plasmid 30(2):170–175. https://doi.org/10.1006/ plas.1993.1047 7. Knudtson KL, Minion FC (1993) Construction of Tn4001lac derivatives to be used as promoter probe vectors in mycoplasmas. Gene 137(2):217–222. https://doi.org/10. 1016/0378-1119(93)90009-r 8. Hahn TW, Mothershed EA, Waldo RH et al (1999) Construction and analysis of a modified Tn4001 conferring chloramphenicol resistance in Mycoplasma pneumoniae. Plasmid 41(2): 120–124. https://doi.org/10.1006/plas. 1998.1387 9. Montero-Blay A, Miravet-Verde S, LluchSenar M et al (2019) SynMyco transposon: engineering transposon vectors for efficient transformation of minimal genomes. DNA Res 26(4):327–339. https://doi.org/10. 1093/dnares/dsz012 10. Pour-El I, Adams C, Minion FC (2002) Construction of mini-Tn4001tet and its use in Mycoplasma gallisepticum. Plasmid 47(2): 129–137. https://doi.org/10.1006/plas. 2001.1558

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11. Reece-Hoyes JS, Walhout AJM (2018) Gateway recombinational cloning. Cold Spring Harb Protoc 2018(1). https://doi.org/10. 1101/pdb.top094912 12. Miyata M, Robinson RC, Uyeda TQP et al (2020) Tree of motility – a proposed history of motility systems in the tree of life. Genes Cells 25(1):6–21. https://doi.org/10.1111/ gtc.12737 13. Miyata M, Hamaguchi T (2016) Integrated information and prospects for gliding mechanism of the pathogenic bacterium Mycoplasma pneumoniae. Front Microbiol 7:960. https:// doi.org/10.3389/fmicb.2016.00960 14. Kenri T, Suzuki M, Sekizuka T et al (2020) Periodic genotype shifts in clinically prevalent Mycoplasma pneumoniae strains in Japan. Front Cell Infect Microbiol 10:385. https://doi. org/10.3389/fcimb.2020.00385 15. Nakane D, Murata K, Kenri T et al (2021) Molecular ruler of the attachment organelle in Mycoplasma pneumoniae. PLoS Pathog 17(6): e1009621. https://doi.org/10.1371/journal. ppat.1009621 16. Nakane D, Kenri T, Matsuo L et al (2015) Systematic structural analyses of attachment organelle in Mycoplasma pneumoniae. PLoS Pathog 11(12):e1005299. https://doi.org/ 10.1371/journal.ppat.1005299 17. Hayflick L (1965) Tissue cultures and mycoplasmas. Tex Rep Biol Med 23(Suppl 1): 285–303 18. Vizarraga D, Kawamoto A, Matsumoto U et al (2020) Immunodominant proteins P1 and P40/P90 from human pathogen Mycoplasma pneumoniae. Nat Commun 21(1):5188. https://doi.org/10.1038/s41467-02018777-y

Chapter 30 Purification and ATPase Activity Measurement of Spiroplasma MreB Daichi Takahashi, Ikuko Fujiwara, and Makoto Miyata Abstract Spiroplasma is a genus of wall-less helical bacteria with swimming motility unrelated to conventional types of bacterial motility machinery, such as flagella and pili. The swimming of Spiroplasma is suggested to be driven by five classes of MreB (MreB1-MreB5), which are members of the actin superfamily. In vitro studies of Spiroplasma MreBs have recently been conducted to evaluate their activities, such as ATPase, which is essential for the polymerization dynamics among classic actin superfamily proteins. In this chapter, we describe methods of purification and Pi release measurement of Spiroplasma MreBs using column chromatography and absorption spectroscopy with the molecular probe, 2-amino-6-mercapto-7-methylpurine riboside (MESG). Of note, the methods described here are applicable to other proteins that possess NTPase activity. Key words Bacterial actin cytoskeleton, E. coli expression system, Recombinant protein, Ni2+-NTA affinity chromatography, Gel filtration, Pi release assay, Absorption spectroscopy

1

Introduction Spiroplasma is one of the genera of small infectious bacteria in the class Mollicutes [1, 2]. Species of this genus have a helical-shaped cell that lacks a cell wall (i.e., the peptidoglycan layer) [3, 4]. As a result, these bacteria do not possess conventional types of bacterial motility machinery, such as flagella and pili. However, Spiroplasma has acquired a unique motility system in its evolutionary path [5]. In fact, these bacteria swim in a viscous liquid by rotating their helical cell body. For cell body rotation, Spiroplasma switches cell helicity and transmits the switch along the cell axis [3, 6, 7]. This swimming is thought to be driven by six cytoskeletal proteins, a fibril that is unique to Spiroplasma and five classes of MreB proteins (MreB1-MreB5) [3, 5, 8–13].

Tohru Minamino et al. (eds.), Bacterial and Archaeal Motility, Methods in Molecular Biology, vol. 2646, https://doi.org/10.1007/978-1-0716-3060-0_30, © The Author(s), under exclusive license to Springer Science+Business Media, LLC, part of Springer Nature 2023

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Fig. 1 Working model of Spiroplasma MreB polymerization dynamics based on a recent publication [16]. ATP-bound MreB molecules polymerize into a filament, while it is not the most stable state as the filament. The MreB molecules hydrolyze ATP to achieve an ADP-Pi bound state. They stochastically release Pi to achieve the ADP bound state and depolymerize. ADP-bound MreB monomers replace the bound nucleotide to ATP and return to the initial phase of the polymerization cycle

MreB is an actin superfamily protein that is conserved in the bacterial kingdom [14]. Upon binding of a nucleotide, such as ATP and GTP, MreB assembles into filamentous structures [15]. Although studies on polymerization dynamics of MreB are limited, our recent study suggested that the ATPase activity of MreB is essential for the polymerization dynamics of MreB, as well as other actin superfamily proteins (Fig. 1) [16, 17]. Although majority of the roles of each class of Spiroplasma MreB remain unclear, previous studies have suggested that the polymerization dynamics of some MreB classes are related to the Spiroplasma swimming [8, 18]. Therefore, in vitro studies of Spiroplasma MreBs have recently been conducted to evaluate their activities, such as ATPase [10, 16, 19]. In this chapter, we describe methods of purification and Pi release measurement of Spiroplasma MreBs, which we have recently applied to MreB3 and MreB5 of S. eriocheiris [16]. Briefly, MreB3 and MreB5 are overexpressed in E. coli as fusions with a 6× histidine tag and are purified by Ni2+-affinity chromatography, followed by gel filtration chromatography. The purified MreBs are polymerized by the addition of ATP. The concentration of released Pi is estimated by absorption spectroscopy with a molecular probe, 2-amino-6-mercapto-7-methylpurine

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riboside (MESG), which shifts the maximum absorption wavelength from 330 to 360 nm by reacting with a Pi molecule [20, 21]. The methods described here are applicable to other proteins that possess NTPase activity.

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Materials All solutions are prepared using Milli-Q water and stored at room temperature, unless otherwise stated. The use of an autoclave is not necessary, if not stated.

2.1

E. coli Strains

1. E. coli BL21 (DE3) or C43 (DE3).

2.2

Plasmid

1. pET15b-based plasmid encoding mreB3 or mreB5 under a control of the T7 promoter (see Note 1).

2.3 Cell Cultivation and Harvesting

1. Toothpick sterilized in an autoclave at 121 °C for 20 min. 2. Glass test tube with an aluminum cap. 3. Luria broth (LB): 1% (w/v) Bacto tryptone, 0.5% (w/v) yeast extract, and 1.0% (w/v) NaCl sterilized in an autoclave at 121 ° C for 20 min. 4. 50 mg/mL ampicillin-sodium (see Notes 2 and 3). 5. Shaker. 6. 2 L glass Erlenmeyer flask with a silicon cap with fine air vents. 7. Spectrophotometer. 8. 1 M isopropyl β-D-1-thiogalactopyranoside (IPTG) (see Note 2). 9. 500 mL bottle. 10. 50 mL tube. 11. 10× phosphate-buffered saline (PBS): 100 mM Na2HPO4, 20 mM KH2PO4, 30 mM KCl, 1.37 M NaCl (see Note 4). 12. Centrifuge for 500 mL bottles and 50 mL tubes. 13. Deep freezer.

2.4

MreB Purification

1. Evaporator. 2. His trap buffer A: 50 mM Tris–HCl, pH 8.0, 50 mM imidazole-HCl, pH 8.0, 300 mM NaCl (see Note 5). 3. His trap buffer B: 50 mM Tris–HCl, pH 8.0, 500 mM imidazole-HCl, pH 8.0, 300 mM NaCl (see Note 5). 4. Gel filtration buffer: 20 mM Tris–HCl, pH 8.0, 300 mM NaCl (see Note 5). 5. Bucket containing ice.

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6. 100 mM phenylmethylsulfonyl fluoride (PMSF) in 100% (v/v) ethanol (see Note 2). 7. Probe sonicator (e.g., Ultrasonic Homogenizer US-600, NISSEI). 8. Centrifuge with temperature stabilizer at 4 °C for 50 mL tubes. 9. Needle-less syringe. 10. Syringe filter unit with 0.45 μm pores. 11. 50 mL tube. 12. 15 mL tube. 13. 96-well plate with fraction volume of 2 mL. 14. Column for Ni2+-NTA affinity chromatography. 15. Column for gel filtration chromatography in which the separation range includes around 40 kDa molecular mass (e.g., HiLoad™ 26/600 Superdex™ 200 pg, Cytiva). 16. Laboratory refrigerator. 17. Fast protein liquid chromatography (FPLC) system (e.g., ¨ KTA pure™ 25, Cytiva). A 18. 12.5% (w/v) SDS-polyacrylamide gel and the gel apparatus for SDS-PAGE. 2.5 Pi Release and Pi Standard Measurements

1. Cellulose dialysis membrane with a 14 kDa cutoff. 2. Microwave oven. 3. Polypropylene tapper. 4. Closure clips for the dialysis membrane. 5. Laboratory refrigerator. 6. Magnetic stirrer. 7. Pre-polymerization buffer: 57.1 mM Tris–HCl, pH 7.5, 285.7 mM KCl. 8. Centrifuge with temperature stabilizer at 4 °C for 50 mL and 1.5 mL tubes. 9. Bucket containing ice. 10. Ultrafiltration unit with a 10 kDa cutoff (e.g., Amicon® Ultra15, Merck). 11. Spectrophotometer. 12. 50 mL tube. 13. 2 mL microtube. 14. 1.5 mL microtube. 15. 0.2 mL microtube with eight strips. 16. 96-well plate with fraction volume of 200 μL and flat and transparent bottoms.

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17. Plate reader with a temperature stabilizer at 25 °C. 18. Polymerization agent (40 mM ATP and 20 mM MgCl2) (see Notes 2 and 6). 19. 50 mM (+/-)-dithiothreitol (DTT) (see Note 2). 20. EnzChek™ Phosphate Assay Kit (Thermo Fisher Scientific) (see Note 7) containing 20× reaction buffer (1.0 M Tris– HCl, pH 7.5 and 20 mM MgCl2), 50 U purine nucleoside phosphorylase (PNP) powder, powder of 6.3 mg MESG (20 μmoles), and phosphate standard solution (50 mM KH2PO4 and 2 mM sodium azide). 21. Block incubator.

3

Methods

3.1 Cell Cultivation and Harvesting

1. Add 5 μL of 50 mg/mL ampicillin-sodium to a 5 mL volume of LB medium in a capped test tube. 2. Pick a colony or frozen stock of E. coli carrying the pET15bbased plasmid encoding either mreB3 or mreB5 with a toothpick and transfer to the LB medium (see Note 1). 3. Shake the culture at 37 °C for 12–18 h (see Note 8). 4. Add 1 mL of overnight culture and 1 mL of 50 mg/mL ampicillin-sodium into 1 L of fresh LB medium in a capped Erlenmeyer flask. 5. Shake the culture at 37 °C and measure its OD600 at different timepoints using a spectrophotometer until a value between 0.4 and 0.6 is achieved (see Note 9). 6. Terminate cultivation when the OD600 of the culture reaches the desired value and add 1 mL of 1 M IPTG to the culture at a final concentration of 1 mM (see Note 10). 7. Shake the culture at 15 °C for 18–24 h. 8. Transfer the culture to 500 mL bottles, adjust the balance, and centrifuge at >6000 × g for 10 min at 4–25 °C. 9. Discard the supernatant and resuspend each pellet in 40 mL of 1× PBS. 10. Transfer each cell suspension to a 50 mL tube and centrifuge using the same conditions described in step 8. 11. Discard the supernatant, resuspend each pellet with several mL of 1× PBS, place cell suspensions in an appropriate number of 50 mL tubes, and centrifuge using the same conditions described in step 8. 12. Discard the supernatant and store the tube(s) at -80 °C until needed.

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3.2 Purification of MreB

1. Resuspend the frozen cell stock in His trap buffer A at a volume of 20–40 mL per cell pellet collected from the 1 L culture. 2. Add 100 mM PMSF at a final concentration of 1 mM (see Note 11). 3. Homogenize the cell with a probe sonicator by repeating several seconds of sonics and several seconds of intervals on ice or ice water (see Note 12). 4. Centrifuge the sample at 12,000 × g for 30 min at 4 °C (see Note 13). 5. Make pass the supernatant through a syringe filter unit with 0.45 μm pores into a 50 mL tube precooled on ice. 6. Load the sample into a Ni2+-NTA affinity chromatography column connected with an FPLC system, remove nonspecifically bound molecules via washing with 10 column volumes of His trap buffer A, and elute MreB molecules bound on the column with 2.6 column volumes of mixed buffer comprising His trap buffers A and B at a ratio of 6:4 (Fig. 2a) (see Notes 14 and 15). 7. Load the sample eluted from the Ni2+-NTA column into a gel filtration chromatography column connected with an FPLC system at 4 °C in a laboratory refrigerator, allow the gel filtration buffer flow through the column, and collect each 2 mL of the eluted solution in 96-well plates placed in the fraction collector (Fig. 2b) (see Notes 14 and 16). 8. Analyze the eluted fractions containing MreB using SDSPAGE (Fig. 2c). 9. Store the plate containing MreB at 4 °C until needed (see Notes 17 and 18).

3.3 Pi Release Measurement of MreB

1. Dissolve 50 U PNP powder in 500 μL Milli-Q water to obtain a 100 U/mL solution (see Note 18). 2. Dissolve 20 μmoles of MESG powder in 20 mL Milli-Q water to make a 1 mM solution and dispense solution into 400 μL aliquots (see Note 2). 3. Place a dialysis membrane in Milli-Q water and boil in a microwave oven to remove the interior and exterior protective materials. 4. Cool the membrane using running Milli-Q water and clip one side of the membrane. 5. Place the sample solution in the membrane and clip the other side of the membrane. 6. Place the sample containing the membrane in a pre-polymerization buffer precooled at 4 °C with a 50–100 times larger volume than the sample (see Note 19).

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Fig. 2 Typical purification results of MreB5 expressed in E. coli C43 (DE3) carrying a fusion of pET-15b and a codon optimized mreB5 gene. (a) An elution pattern of Ni2+-NTA affinity chromatography traced by the absorbance at 280 nm. The flowthrough (1), pre-wash with buffer A (2), elution (3), and final-wash with buffer B (4) fractions are indicated by each corresponding step number. (b) An elution pattern of gel filtration chromatography traced by the absorbance at 280 nm. The triangle indicates the elution peak of MreB5. The elution volumes of bovine γ-globulin (158 kDa), chicken ovalbumin (44 kDa), and horse myoglobin (17 kDa) are plotted as diamonds over the log of their molecular weight. The estimated molecular mass of MreB5 is 42 kDa, which aligns with that estimated from the amino acid sequence (see Note 20), indicating that it is purified as a monomer. (C) SDS-PAGE analysis of the purification procedure. Lanes 1, 2, 3, 4, and 5 contain the whole cell lysate (step 3), the soluble fraction of the lysate (step 4), the insoluble fraction of the lysate (step 4), the flowthrough fraction of Ni2+-NTA affinity chromatography (step 6), and the eluted fraction of Ni2+-NTA affinity chromatography (step 6), respectively. The right six lanes indicate the elution peak of the gel filtration chromatography on which the elution volumes are indicated. Protein size standards are shown in lane M with the molecular masses of each band on the left side

7. Stir the buffer at 4 °C for more than 8 h. 8. Salvage the membrane, remove the clip from one side, and transfer the sample to a 50 mL tube precooled on ice. 9. Measure the concentration of the sample with a spectrophotometer (the concentration should be 2.86-fold higher than the desired final concentration) (see Note 20).

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10. If the concentration of the sample is lower than the desired concentration, concentrate the sample with ultrafiltration unit. 11. Transfer the sample to a 1.5 mL microtube precooled on ice and centrifuge at 20,000 × g for 10 min at 4 °C to remove aggregates. 12. Transfer the supernatant to another 1.5 mL microtube precooled on ice. 13. Measure the concentration of the sample with a spectrophotometer and dilute with a pre-polymerization buffer precooled on ice to obtain a concentration 2.86-fold higher than the desired final concentration on ice (see Note 20). 14. Prepare two 1.5 mL microtubes precooled on ice and mix solutions to obtain sample premixes on ice as follows (for three aliquots each) (Fig. 3a). Mix 280 μL sample solution, 192 μL Milli-Q water, and 80 μL of 50 mM DTT, and add 8 μL of 100 U/mL PNP (for samples with MESG) or Milli-Q water (for samples without MESG). 15. Prepare two 1.5 mL microtubes precooled on ice and mix solutions to obtain background premixes on ice as follows (for three aliquots each) (Fig. 3a). Mix 70 μL of pre-polymerization buffer, 48 μL of Milli-Q water, and 20 μL of 50 mM DTT, and add 2 μL of 100 U/mL PNP (for background with MESG) or Milli-Q water (for background without MESG). 16. Prepare a 0.2 mL microtube with eight strips and mix solutions to obtain reaction premixes as follows (Fig. 3a). Mix 11 μL 20× reaction buffer and 11 μL polymerization agent in each strip, and add 44 μL of 1 mM MESG (for strips 1–3 and 7) or MilliQ water (for strips 4–6 and 8) (see Note 21). 17. Dispense the sample and background premixes prepared in steps 14 and 15 into 140 μL aliquots in a 96-well plate in the order indicated in Fig. 3a. 18. Pause until the dew condensation on the bottoms of the plate disappears at room temperature. 19. Add 60 μL of the reaction premixes prepared in step 16 to the aliquots as indicated in Fig. 3a (see Note 22). 20. Place the plate in a plate reader and start a time-course measurement of the absorbances at 360 nm. 3.4 Acquisition of the Pi Standard

1. Prepare nine 1.5 mL microtubes and a 2 mL microtube. 2. Add 200 μL Milli-Q water into eight 1.5 mL microtubes. 3. Mix 498 μL Milli-Q water and 2 μL phosphate standard solution in the remaining empty 1.5 mL microtube to prepare 200 μM KH2PO4 solution.

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Fig. 3 Pi release measurement of MreB5. (a) Scheme of the premix preparation in a 96-well plate and a 0.2 mL microtube with eight strips. In a 96-well plate, the sample premixes with and without PNP and the background premixes with and without PNP are dispensed in lines 1–3, 4–6, 7, and 8, respectively. In a 0.2 mL microtube with eight strips, the reaction premix with MESG is prepared into strips 1, 2, 3, and 7. The reaction premix without MESG is prepared into the other strips. As a result, the sample solutions with and without MESG-PNP and the background solutions with and without MESG-PNP are prepared in lines 1–3, 4–6, 7, and 8, respectively. (b) Pi standard curve. The function of the linear fit and the coefficient of determination are indicated in the graph. (c) Pi releases of 3 μM MreB5 measured from three independent samples

4. Add 200 μL of 200 μM KH2PO4 solution to a tube containing 200 μL Milli-Q water to prepare a twofold dilution of the solution. 5. Add 200 μL of the mixture to another tube containing 200 μL Milli-Q water. 6. Repeat step 5 to prepare eight tubes containing KH2PO4 with the double diluted concentration series. A total of eight tubes containing KH2PO4 and a tube containing Milli-Q should be prepared (see Note 23). 7. Discard the tube used in step 3 as well as 200 μL of the final mix. A final volume of 200 μL is required. 8. Mix 912 μL Milli-Q water, 190 μL of 20× reaction buffer, 760 μL of 1 mM MESG, and 38 μL of 100 U/mL PNP in a 2 mL microtube to prepare a reaction premix (see Note 21).

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9. Add 200 μL of the reaction premix to each tube containing 200 μL KH2PO4 solution and Milli-Q water. 10. Incubate the solutions at 25 °C for 30 min. 11. Prepare 200 μL of aliquots of the solution in a 96-well plate. 12. Place the plate in a plate reader and measure the absorbances at 360 nm. 3.5 Construction of Pi Standard Curve

1. Import the dataset acquired in Subheading 3.4 into a spreadsheet. 2. Average the absorbances derived from the solutions at the same KH2PO4 concentration. 3. Subtract the absorbance of 0 μM KH2PO4 solution from the absorbance of all solutions. 4. Plot the final KH2PO4 concentrations relative to the values obtained in step 3 (Fig. 3b) (see Note 23). 5. Calculate the slope of the linear fit.

3.6 Construction of Pi Release Curve of MreB

1. Import the dataset acquired in Subheading 3.3 into a spreadsheet. 2. Subtract the time-course absorbances of the background with MESG from those of the MreB solutions with MESG. 3. Subtract the time-course absorbances of the background without MESG from those of the MreB solutions without MESG. 4. Subtract one of the set of values of the time-course trace without MESG, estimated in step 3, from one of those with MESG, estimated in step 2. 5. Subtract the value at time 0 from all values over the trace. 6. Multiply the slope estimated in step 5 of Subheading 3.5 by all values over the trace. The calculated values are the estimated Pi concentrations of the sample. 7. Plot the Pi concentrations estimated in step 6 over time. The rate constant for Pi release of MreB can be estimated from the slope of the plot (see Note 24). 8. Repeat steps 4–7 for all data.

4

Notes 1. MreB3 and MreB5 are sometimes not expressed using pET-15b. When the expression levels are not well controlled, other vectors, such as pCold-I (Takara Bio), can be used. 2. Store at -20 °C and do not heat the solution after thawing. To avoid degradation of the compounds, do not refreeze the thawed solutions.

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3. Prepare the solution with 50% (v/v) ethanol and filter with a sterilized syringe filter unit with 0.22 μm pores. 4. Do not store at 1× dilution to avoid stock solution degradation. The solution should be diluted to 1× with Milli-Q water prior to use. 5. Make pass all solutions for column chromatography through a filter with 0.45 μm pores and degas the solutions with an evaporator. Otherwise, the flow channels of FPLC systems and the resin in the columns will be damaged. 6. MgCl2 solution can be stored separately at room temperature. Immediately after the ATP powder is dissolved in Milli-Q water, adjust the pH to approximately 7 using NaOH or KOH as the ATP solution is acidic. 7. Other kits containing MESG and PNP, such as ATPase Kinetic ELIPA Assay Kit (Cytoskeleton, Inc.) and PhosphoWorks™ Colorimetric MESG Phosphate Assay Kit *UV absorption* (COSMO BIO CO., LTD.) can be used although some steps should be modified due to the differences of some contents. 8. The overnight culture can be stored at -80 °C for several years by mixing with autoclaved 50% (v/v) glycerol at a ratio of 5:3 (culture:glycerol). 9. Adjust the baseline with LB medium. 10. When the pCold vector is used, cool the culture on ice for several seconds, and incubate at 15 °C for 30 min without shaking before adding IPTG. This process is necessary for IPTG induction using pCold vectors. 11. PMSF should be added immediately before homogenization as the half-life of PMSF in the aqueous phase is 30 min. 12. An interval is required as consecutive sonication heats the tip of the probe and damages the sample. 13. Ultra-centrifugation at 100,000 × g for 30 min at 4 °C is also applicable. If performed, filtration of the supernatant can be skipped. 14. Prior to loading the sample, equilibrate columns for Ni2+-NTA affinity and gel filtration chromatography with His trap buffer A and gel filtration buffer, respectively. After the use, replace the flow channel of the FPLC and the columns with degassed Milli-Q water. 15. The sample application and buffer flow to some Ni2+-NTA columns can be performed manually using a needle-less syringe. To avoid the contamination of molecules resting in the column, wash the column with five column volumes of His trap buffer B before the equilibration with His trap buffer A and after the elution of the sample.

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16. Restrict the sample volume for gel filtration chromatography to 4% of the column bed volume to avoid broadening the elution peaks. When the sample volume is greater than the limit, the sample should be concentrated by ultrafiltration. 17. The first half of the MreB5 elution peak is prone to aggregation. However, the MreB5 molecules in the supernatant display the same activity as those in the second half of the elution peak. 18. The solution can be stored at 4 °C for 2–3 months. Do not heat. 19. If microbubbles affect the measurement, the prepolymerization buffer should be degassed before this step. 20. If MreB3 and MreB5 are fused with a histidine tag derived from pET-15b, their estimated absorption coefficients and molecular masses are 0.474 and 0.578 mg/mL/cm at 280 nm, respectively, and 40,431.98 and 40,677.98 Da, respectively. These parameters can be estimated from the amino acid sequences using the ProtParam server (https:// web.expasy.org/protparam/). 21. MESG should be added immediately before the initiation of the reaction because the half-life of MESG in the aqueous phase is 4 h at room temperature. 22. The final buffer composition is 40 mM Tris–HCl, pH 7.5; 100 mM KCl; 5 mM DTT; 2 mM MgCl2; 2 mM ATP; 0.2 mM MESG; and 1 U PNP. 23. KH2PO4 concentrations in the dilution series are 100, 50, 25, 12.5, 6.25, 3.13, 1.56, and 0 μM. Of note, the final concentrations of KH2PO4 are two times lower than those resulting from the addition of the reaction premix. 24. If the curve is not linear, estimate the rate from the initial slope.

Acknowledgments This study was supported by Grants-in-Aid for Scientific Research (A and C) (MEXT KAKENHI, Grant Numbers JP17H01544 to MM and JP20K06591 to IF), JST CREST (Grant Number JPMJCR19S5 to MM), the Research Foundation of Opto-Science and Technology to IF, and the Osaka City University (OCU) Strategic Research Grant 2019 to IF. DT is a recipient of the Research Fellowship of the Japan Society for the Promotion of Science (22J10345).

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References 1. Paredes JC, Herren JK, Schu¨pfer F et al (2015) Genome sequence of the Drosophila melanogaster male-killing Spiroplasma strain MSRO endosymbiont. mBio 6(2):e02437–02414 2. Gasparich GE (2002) Spiroplasmas: evolution, adaptation and diversity. Front Biosci 7:d619– d640 3. Liu P, Zheng H, Meng Q et al (2017) Chemotaxis without conventional two-component system, based on cell polarity and aerobic conditions in helicity-switching swimming of Spiroplasma eriocheiris. Front Microbiol 8:58 4. Terahara N, Tulum I, Miyata M (2017) Transformation of crustacean pathogenic bacterium Spiroplasma eriocheiris and expression of yellow fluorescent protein. Biochem Biophys Res Commun 487(3):488–493 5. Miyata M, Robinson RC, Uyeda TQP et al (2020) Tree of motility – a proposed history of motility systems in the tree of life. Genes Cells 25(1):6–21 6. Nakane D, Ito T, Nishizaka T (2020) Coexistence of two chiral helices produces kink translation in Spiroplasma swimming. J Bacteriol 202(8):e00735–e00719 7. Shaevitz JW, Lee JY, Fletcher DA (2005) Spiroplasma swim by a processive change in body helicity. Cell 122(6):941–945 8. Kiyama H, Kakizawa S, Sasajima Y et al (2022) Reconstitution of minimal motility system based on Spiroplasma swimming by two bacterial actins in a synthetic minimal bacterium. Sci Adv 8(48):eabo7490 9. Takahashi D, Fujiwara I, Miyata M (2020) Phylogenetic origin and sequence features of MreB from the wall-less swimming bacteria Spiroplasma. Biochem Biophys Res Commun 533(4):638–644 10. Harne S, Duret S, Pande V et al (2020) MreB5 is a determinant of rod-to-helical transition in the cell-wall-less bacterium Spiroplasma. Curr Biol 30(23):4753–4762.e4757 11. Ku C, Lo WS, Kuo CH (2014) Molecular evolution of the actin-like MreB protein gene

family in wall-less bacteria. Biochem Biophys Res Commun 446(4):927–932 12. Trachtenberg S, Dorward LM, Speransky VV et al (2008) Structure of the cytoskeleton of Spiroplasma melliferum BC3 and its interactions with the cell membrane. J Mol Biol 378(4):778–789 13. Ku¨rner J, Frangakis AS, Baumeister W (2005) Cryo-electron tomography reveals the cytoskeletal structure of Spiroplasma melliferum. Science 307(5708):436–438 14. Shi H, Bratton BP, Gitai Z et al (2018) How to build a bacterial cell: MreB as the foreman of E. coli construction. Cell 172(6):1294–1305 15. Wagstaff J, Lo¨we J (2018) Prokaryotic cytoskeletons: protein filaments organizing small cells. Nat Rev Microbiol 16(4):187–201 16. Takahashi D, Fujiwara I, Sasajima Y et al (2022) ATP-dependent polymerization dynamics of bacterial actin proteins involved in Spiroplasma swimming. Open Biol 12(10): 220083 17. Wegner A (1976) Head to tail polymerization of actin. J Mol Biol 108(1):139–150 18. Masson F, Pierrat X, Lemaitre B et al (2021) The wall-less bacterium Spiroplasma poulsonii builds a polymeric cytoskeleton composed of interacting MreB isoforms. iScience 24(12): 103458 19. Pande V, Mitra N, Bagde SR et al (2022) Filament organization of the bacterial actin MreB is dependent on the nucleotide state. J Cell Biol 221(5):e202106092 20. Blanchoin L, Pollard TD (1999) Mechanism of interaction of Acanthamoeba actophorin (ADF/cofilin) with actin filaments. J Biol Chem 274(22):15538–15546 21. Webb MR (1992) A continuous spectrophotometric assay for inorganic-phosphate and for measuring phosphate release kinetics in biological-systems. Proc Natl Acad Sci U S A 89(11):4884–4887

Chapter 31 Swimming Motility Assays of Spiroplasma Daisuke Nakane Abstract Spiroplasma swim in liquids without the use of the bacterial flagella. This small helical bacterium propels itself by generating kinks that travel down the cell body. The kink translation is unidirectional, from the leading pole to the lagging pole, during cell swimming in viscous environments. This protocol describes a swimming motility assay of Spiroplasma eriocheiris for visualizing kink translations of the absolute handedness of the body helix with optical microscopy. Key words Cell motility, Helical shape, Cell polarity, Mollicutes, Optical microscopy

1

Introduction Spiroplasma, small wall-less bacteria, swim without flagellar rotation in viscous environments though the mechanism is currently not well understood [1–3]. In contrast to the rotary motion of bacterial flagella, the propulsion of Spiroplasma is generated be waves of helical transformation. They possess both left-handed helix (LH) and right-handed helix (RH) chiral shapes in a single cell [1, 4]. To maintain the bistable helices, the cell bends at the boundary of the helicity switch at a specific angle, which is known as a kink [1, 5, 6]; the kink translation is unidirectional along the cell body during cell swimming, from the leading pole to the lagging pole [1, 4] (Fig. 1a–c). This is possibly accompanied by dual rotation of two helices in opposite directions, thereby generating efficient propulsion force and cancelling torque around the helical axis [7–9]. The characteristic cell shape has been defined by the bacterial cytoskeletal protein MreB and the Spiroplasma-specific protein fibril, with the filaments of these proteins located at the innermost line of the body helix [10–12]. The Spiroplasma genome contains several MreB homologues [13–15], and interactions between MreB isoforms induce dynamic cell morphological changes during swimming motility [16, 17]. Synthetic biology

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Fig. 1 Processive change of body helicity in Spiroplasma eriocheiris during swimming. (a) Schematic of the cell morphology. Left: Kinked Spiroplasma cell. The cell moves in the direction of the white arrow. A kink appears at the front end, and the translation is unidirectional toward the rear end, presumably to generate a propulsion force in the same direction by the dual rotation of the RH and LH. Middle: Kink at the boundary of the two chiral helices. The helical cell shape was designated based on the slanted portions of the cell. The two helical axes are connected with a bend called kink. Right: Geometry of the kink. The bend angle at the kink was related to the pitch angles of the RH and LH. (b) Sequential phase-contrast microscopy images of a single cell near the glass surface. The images were captured at an interval of 1/30 s. The cell shapes of the RH and LH were based on the slanted portions of the cell. The schematic and images in panel AB are shown as mirror images captured by an inverted microscope. (c) Kink diagram, a graphical representation of the processive change in body helicity. The distance from the rear end to the kink is plotted at each time point during swimming. (d) Sample chamber. Two pieces of double-sided tape are used to assemble two coverslips. To replace the solution in the chamber, a piece of filter paper is placed at one side of the tunnel in the chamber, and add the new solution such as cell suspension from the other side

has revealed that the coexistence of MreB isoforms is sufficient for the spontaneous self-organization of the helical morphology and the generation of kinks, but the kink propagation is bidirectional [18, 19]. Given that the native Spiroplasma cell has a dumbbell-like structure at the front end [20], an additional factor is required for unidirectional kink propagation to achieve directional swimming motility in vivo [3]. The unidirectional waves of helical transformation are characteristic features for the mechanism of Spiroplasma swimming. However, it seems challenging to determine the absolute handedness of the body helix because the position of the focal plane is ambiguity during the swimming motility. In Spiroplasma eriocheiris, a pathogen of Chinese mitten crab, the cell morphology is

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Fig. 2 Handedness of the cell body in a stopped cell. (a) Cell images under phase-contrast microscopy. Cell motility was stopped by the addition of CCCP (left) and strong light irradiation (right). Light irradiation was applied in the presence of FM4-64. The slanted portions of the cell are indicated by the arrows. (b) Absolute handedness of the body helicity determined by the cell image. Left upper: The cell movement was observed in the chamber near the bottom side of the glass surface. The focal plane was fixed slightly below the cell, as indicated by the black arrow. Right: Cell images captured as mirror images under an inverted microscope. Left lower: The orientation of the original specimen and that projected back to the camera

fixed to be RH and LH by adding a conventional uncoupler, carbonyl cyanide 3-chlorophenylhydrazone (CCCP) and a strong light irradiation, respectively [4]. Additionally, the cells preferentially swim at the same image plane near the surface of an observation when the chamber is pre-treated with bovine serum albumin (BSA) [4]. By the combination of these methods, the cell helicity can be determined by the direction of the slanted portions of the cells relative to the helical axis even under the conventional phasecontrast microscopy (Fig. 2a). Here, I describe a method to visualize the swimming motility of S. eriocheiris in a viscous environment by phase-contrast microscopy [4] (Fig. 1b). In addition, the method to observe the unidirectional waves of helical transformation with the absolute handedness of body helix is described (Fig. 1c).

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Materials Prepare all solutions using ultrapure water and analytical grade reagents. Sterilize them (autoclave for 20 min at 121 °C) prior to use, and store all reagents at 4 °C (unless indicated otherwise).

2.1 Growth Medium and Bacterial Strains

1. R2 liquid medium to grow Spiroplasma eriocheiris TDA-040725-5 cells for the motility assay [21] (see Note 1): 2.5% (w/v) heart infusion broth, 8% (w/v) sucrose, 15% (v/v) newborn calf serum (see Note 2). Add newborn calf serum after autoclave.

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2. Incubator at 30 °C. 3. Tissue culture flask (growth surface 25 cm2). 4. Spectrophotometer to measure an optical density at a wavelength of 600 nm. 2.2 Optical Microscopy and Data Analysis

1. Inverted microscope (e.g., IX73, Olympus). 2. Objective lens for phase-contrast UPLSAPO100 × O2PH, Olympus).

microscopy

(e.g.,

3. Condenser for phase-contrast microscopy. 4. CMOS camera (e.g., Zyla 4.2, Andor or DMK33U174, Imaging Source). 5. Optical filter (e.g., excitation; FF01-531/40, and dichroic mirror; FF593-Di03, Semrock). 6. Epi-illuminator with a mercury lamp (e.g., U-HGLGPS, Olympus). 7. Software for image analysis such as ImageJ (http://rsb.info. nih.gov/ij/). 2.3 Glass Chamber Assembly

1. Coverslip: 22 × 40 mm. 2. Coverslip: 18 × 18 mm. 3. Double-sided tape. 4. Filter paper. 5. Nail polish.

2.4

Motility Assay

1. MC: 2% (w/v) methylcellulose (viscosity 4000 cP) in water. Dissolve 2 g MC in water to a volume of 100 mL on a hot plate stirrer. After autoclaving, cool the mixture slowly on a stirrer overnight to a temperature of 4 °C. Use this as stock solution. 2. Motility buffer (MB): 75 mM Na2HPO4/NH2PO4 (pH 7.3), 68 mM NaCl, 235 mM sucrose, and 0.25–0.5% (w/v) MC. 3. 20% (w/v) BSA: 20% (w/v) bovine serum albumin (BSA) in MB.

2.5 Stopping Cell Motility

1. CCCP: 500 μM carbonyl cyanide 3-chlorophenylhydrazone in MB (see Note 3). 2. FM4-64: N-(3-triethylammoniumpropyl)-450 μM (6-(4-(diethylamino) phenyl) hexatrienyl) pyridinium dibromide in MB (see Note 4).

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Methods Carry out all procedures at room temperature unless otherwise specified.

3.1

Cell Preparation

1. Add 10 mL of the R2 medium to a tissue culture flask. 2. Inoculate 0.1 mL of the frozen stock of Spiroplasma eriocheiris in the tissue culture flask (see Note 5). 3. Incubate the flask at 30 °C until OD600 of the cell culture reachs 0.06–0.10 (see Note 6).

3.2 Setting for Cell Observation

1. Assemble a sample chamber with double-sided tape to separate the coverslips (Fig. 1d) (see Note 7). 2. Pour 20% (w/v) BSA to the sample chamber for surface coating. 3. Place a piece of filter paper on one side of the tunnel in the chamber, and add MB from the other side to replace the solution in the chamber (Fig. 1d). Repeat this process to until all unbound BSA completely removed. 4. Mix the cell culture with MC to until it reaches a final concentration of 0.25–0.50% (w/v). 5. Place a piece of filter paper on one side of the tunnel in the chamber, and pour the cell suspension into the sample chamber to replace the solution. 6. Seal both the ends of the chamber with nail polish to ensure that the sample do not dry out.

3.3 Cell Observations with Phase-Contrast Microscopy

1. Place a sample chamber on the microscope stage of a phasecontrast microscope. 2. Focus on cells that move over the surfaces on the lower side of the glass chamber. 3. Project the cell images to a camera (see Note 8). 4. Keep the focal plane slightly defocused below the cell to emphasize the direction of the slanted portions of the righthanded (RH) and left-handed (LH) helices (Fig. 1b) (see Note 9). 5. Record the sequential images of the cells at an exposure time of 0.01 s (see Note 10). 6. Observe the cells within 20 min.

3.4 Measuring the Kink Position After Recording

1. Open the sequential images of the swimming cell by software such as ImageJ.

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2. Track the position of the kink, and measure the distance between the tip and the kink (see Note 11). 3. Plot the vertical axis as the length, and the horizonal axis as the observation time (Fig. 1c) (see Note 12). 3.5 Inhibition of Cell Motility with CCCP

1. Place a piece of filter paper on one side of the tunnel in the chamber, and add the CCCP to the sample chamber after steps 1–9 in Subheading 3.2 are completed (see Note 13). 2. Seal the both ends of the chamber with nail polish to ensure that the sample did not dry out. 3. Focus on the cells moving over the surfaces on the lower side of glass, and keep the focal plane slightly defocused below the cell to emphasize the direction of the slanted portions of the righthanded helix (RH) (Fig. 2a).

3.6 Inhibition of Cell Motility with Light Irradiation

1. Place a piece of filter paper on one side of the tunnel in the chamber, and add the FM4-64 to the sample chamber after steps 1–9 in Subheading 3.2 are completed. 2. Seal the both ends of the chamber with nail polish to ensure that the sample did not dry out. 3. Apply a strong light at 2 × 107 W/m2 with an epi-illuminator built into the microscope (see Note 14). 4. Focus on the cells moving over the surfaces on the lower side of glass, and keep the focal plane slightly defocused below the cell to emphasize the direction of the slanted portions of the lefthanded helix (LH) (Fig. 2a).

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Notes 1. The Spiroplasma eriocheiris TDA-040725-5 strain was originally isolated from the Chinese mitten crab Eriocheir sinensis [21], and the strain can be obtained from the Institute of DSMZ-German Collection of Microorganisms and Cell Cultures (https://www.dsmz.de/) as DSM No. 21848. 2. The serum is inactivated in a water bath at 56 °C for 30 min. 3. Dissolve CCCP in dimethyl sulfoxide (DMSO) at 5 mM and dilute it to 500 μM in MB before use. The CCCP concentration described here is ten times higher than that generally used to inhibit the proton motive force (PMF) in flagellar rotation [4, 22]. There is no direct evidence that the PMF drives cell motility in Spiroplasma. 4. Dissolve FM4-64 in DMSO at 5 mM and dilute it to 50 μM in MB before use. The FM4-64 concentration described here is ten times higher than that generally used for membrane

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staining. The high concentration of FM4–64 is effective for inhibiting cell motility with light adsorption. 5. Frozen stock is prepared by the growing a cell culture to an OD600 of 0.1. The culture is transferred to a microtube and is stored at -80 °C. The serum in the medium stabilizes the frozen Spiroplasma like a glycerol stock for bacteria. 6. Use the cells in the late exponential phase. The cell motility is not clear during the stationary phase. The doubling time of Spiroplasma eriocheiris is approximately 5 h under optimal growth conditions. 7. The height and width are approximately 100 μm and 5 mm, respectively, resulting in a tunnel space volume of 10 μL. 8. This is based on observations made with an inverted microscope, which has a mirror between the objective lens and tube lens for projecting images to a camera, which is located at the side port of the microscope (Fig. 2b). 9. In the mirror image, the LH and RH of the cell body is designated based on the slanted portions of the cell image from the lower left to upper right sides and the lower right to upper left sides, respectively [23]. 10. The helical cell shape is not clearly captured at longer exposure times. 11. The helical axis of the cell bends at the kink. The angle is approximately 120°, which is estimated as [180° (θRH + θLH)], where θRH and θLH are the pitch angles of the RH and LH, respectively [1, 4, 5]. 12. The analysis was originally performed in Spiroplasma melliferum [1], and it was slightly modified for Spiroplasma eriocheiris to highlight the directional wave of the kink by displaying the LH and RH regions in different colors [4]. The speed of the kink has a similar distribution between 10 and 15 μm/s in the three related species S. melliferum, S. citri, and S. eriocheiris [1, 4, 24]. 13. Cell motility is completely halted within a few seconds, and the kink no longer appears, which is presumably due to the inhibition of the PMF. The handedness of the body helicity is fixed to RH in Spiroplasma eriocheiris [4]. The S. melliferum cells also stop generating kinks when energy is not supplied [25]. 14. Cell motility is completely stopped after light irradiation, and the kink no longer appears [4]. Strong light irradiation of the fluorescent dye presumably produces reactive oxygen species (ROS) [26, 27], which inhibits kink formation and the propagation of the helicity switch. The handedness of the body helicity is fixed to LH in Spiroplasma eriocheiris [4]. The inhibitory effect of the ROS on kink formation and propagation has also been reported in S. melliferum [28].

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Acknowledgments This study was supported in part by KAKENHI (16H06230, 20H05543, 21 K07020, 22H05066) to DN. References 1. Shaevitz JW, Lee JY, Fletcher DA (2005) Spiroplasma swim by a processive change in body helicity. Cell 122:941–945. https://doi.org/ 10.1016/j.cell.2005.07.004 2. Miyata M, Robinson RC, Uyeda TQP et al (2020) Tree of motility – a proposed history of motility systems in the tree of life. Genes Cells 25:6–21. https://doi.org/10.1111/gtc. 12737 3. Sasajima Y, Miyata M (2021) Prospects for the mechanism of Spiroplasma swimming. Front Microbiol 12:706426. https://doi.org/10. 3389/fmicb.2021.706426 4. Nakane D, Ito T, Nishizaka T (2020) Coexistence of two chiral helices produces kink translation in Spiroplasma swimming. J Bacteriol 202:e00735–e00719. https://doi.org/10. 1128/JB.00735-19 5. Goldstein RE, Goriely A, Huber G et al (2000) Bistable helices. Phys Rev Lett 84:1631–1634. https://doi.org/10.1103/PhysRevLett.84. 1631 6. Wolgemuth CW, Charon NW (2005) The kinky propulsion of Spiroplasma. Cell 122: 827–828. https://doi.org/10.1016/j.cell. 2005.09.003 7. Wada H, Netz RR (2007) Model for selfpropulsive helical filaments: kink-pair propagation. Phys Rev Lett 99:108102. https://doi. org/10.1103/PhysRevLett.99.108102 8. Wada H, Netz RR (2009) Hydrodynamics of helical-shaped bacterial motility. Phys Rev E 80:021921. https://doi.org/10.1103/ PhysRevE.80.021921 9. Yang J, Wolgemuth CW, Huber G (2009) Kinematics of the swimming of Spiroplasma. Phys Rev Lett 102:218102. https://doi.org/ 10.1103/PhysRevLett.102.218102 10. Trachtenberg S, Gilad R (2001) A bacterial linear motor: cellular and molecular organization of the contractile cytoskeleton of the helical bacterium Spiroplasma melliferum BC3. Mol Microbiol 41:827–848. https://doi.org/ 10.1046/j.1365-2958.2001.02527.x 11. Kurner J, Frangakis AS, Baumeister W (2005) Cryo-electron tomography reveals the cytoskeletal structure of Spiroplasma melliferum.

Science 307:436–438. https://doi.org/10. 1126/science.1104031 12. Sasajima Y, Kato T, Miyata T et al (2022) Isolation and structure of fibril protein, a major component of the internal ribbon for Spiroplasma swimming. Front Microbiol 13: 1004601. https://doi.org/10.3389/fmicb. 2022.1004601 13. Ku C, Lo W-S, Kuo C-H (2014) Molecular evolution of the actin-like MreB protein gene family in wall-less bacteria. Biochem Biophys Res Commun 446:927–932. https://doi. org/10.1016/j.bbrc.2014.03.039 14. Takahashi D, Fujiwara I, Miyata M (2020) Phylogenetic origin and sequence features of MreB from the wall-less swimming bacteria Spiroplasma. Biochem Biophys Res Commun 533:638–644. https://doi.org/10.1016/j. bbrc.2020.09.060 15. Takahashi D, Fujiwara I, Sasajima Y, et al (2022) Open Biol 12:220083. https://doi. org/10.1098/rsob.220083 16. Harne S, Duret S, Pande V et al (2020) MreB5 is a determinant of rod-to-helical transition in the cell-wall-less bacterium Spiroplasma. Curr Biol 30:4753–4762.e4757. https://doi.org/ 10.1016/j.cub.2020.08.093 17. Masson F, Pierrat X, Lemaitre B et al (2021) The wall-less bacterium Spiroplasma poulsonii builds a polymeric cytoskeleton composed of interacting MreB isoforms. iScience 24: 103458. https://doi.org/10.1016/j.isci. 2021.103458 18. Kiyama H, Kakizawa S, Sasajima Y et al (2022) Reconstitution of a minimal motility system based on Spiroplasma swimming by two bacterial actins in a synthetic minimal bacterium. Sci Adv 8:eabo7490. https://doi.org/10.1126/ sciadv.abo7490 19. Lartigue C, Lambert B, Rideau F et al (2022) Cytoskeletal components can turn wall-less spherical bacteria into kinking helices. Nat Commun 13:6930. https://doi.org/10. 1038/s41467-022-34478-0 20. Liu P, Zheng H, Meng Q et al (2017) Chemotaxis without conventional two-component system, based on cell polarity and aerobic conditions in helicity-switching swimming of

Swimming Motility Assays of Spiroplasma Spiroplasma eriocheiris. Front Microbiol 8:58. https://doi.org/10.3389/fmicb.2017.00058 21. Wang W, Gu W, Gasparich GE et al (2011) Spiroplasma eriocheiris sp. nov., associated with mortality in the Chinese mitten crab, Eriocheir sinensis. Int J Syst Evol Microbiol 61: 703–708. https://doi.org/10.1099/ijs.0. 020529-0 22. Manson MD, Tedesco P, Berg HC et al (1977) A protonmotive force drives bacterial flagella. Proc Natl Acad Sci U S A 74:3060–3064. https://doi.org/10.1073/pnas.74.7.3060 23. Shimada K, Kamiya R, Asakura S (1975) Lefthanded to right-handed helix conversion in Salmonella flagella. Nature 254:332–334. https://doi.org/10.1038/254332a0 24. Boudet JF, Mathelie´-Guinlet M, Vilquin A et al (2018) Large variability in the motility of spiroplasmas in media of different viscosities. Sci Rep 8:17138. https://doi.org/10.1038/ s41598-018-35326-2

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Chapter 32 Motility Assays of Chloroflexus Shin Haruta, Hinata Kakuhama, Shun-ichi Fukushima, and Sho Morohoshi Abstract Chloroflexus is a thermophilic, filamentous, gliding bacterium. Its multicellular filaments of several hundred micrometer length move straightforward at a speed of approximately 1–3 μm/s and occasionally reverse the moving direction. In liquid media, filaments glide on each other to form cell aggregates without tight adhesion. The molecular machinery on the cell surface that forces the gliding movement has not yet been identified. Here, we describe the cultivation methods to characterize the gliding motility of Chlroflexus and the microscopic assays to determine its gliding speed, reversal frequency, and cell-surface movements. Key words Gliding, Multicellular filament, Anoxygenic phototroph, Thermophile, Cell aggregation

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Introduction The genus Chloroflexus is a phylogenetic group of thermophilic, anoxygenic photosynthetic bacteria in the deeply branched phylum, Chloroflexota [1, 2], and is considered an ancient photosynthetic organism [3]. Chloroflexus can grow photohetero- and photoauto-trophically under anaerobic conditions in the light and chemohetero- and chemoauto-trophically under aerobic conditions in the dark [1, 4, 5]. Bacteria in the genus Chloroflexus are widely distributed in slightly alkaline geothermal springs to frequently form microbial mats [6, 7]. Chloroflexus cells form unbranched multicellular filaments (up to several hundred micrometer long) and move directly along the long axis of the filament via non-flagellated gliding motion [7, 8]. Chloroflexus aggregans shows faster movement at speeds of 1–3 μm/s at 55
C than other Chloroflexus species [9, 10]. C. aggregans forms cell aggregates in liquid media by gliding on other filaments without tight adhesion between filaments [10]. The cell surface movement for gliding motility has been detected [11]. However, the gliding machinery has not been observed and

Tohru Minamino et al. (eds.), Bacterial and Archaeal Motility, Methods in Molecular Biology, vol. 2646, https://doi.org/10.1007/978-1-0716-3060-0_32, © The Author(s), under exclusive license to Springer Science+Business Media, LLC, part of Springer Nature 2023

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genes related to the gliding machinery have not been identified in Chloroflexus [12]. In this chapter, we introduce a variety of methods to characterize the motility of this thermophilic filamentous bacterium with main focus on the fast-gliding species C. aggregans.

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Materials

2.1 Cultivation of Chloroflexus

1. Strains: Chloroflexus aggregans DSM 9485T (¼MD-66T) and NBF [13] (see Note 1). 2. Liquid media: PE medium (per liter, pH 7.5) [9], 0.5 g (NH4)2SO4, 0.5 g Na2S2O3, 0.38 g KH2PO4, 0.39 g K2HPO4, 0.5 g sodium glutamate, 0.5 g disodium succinate hexahydrate, 0.5 g sodium acetate, 0.5 g yeast extract, 0.5 g casamino acids, 1 mL of vitamin mixture, and 5 mL of trace mineral solution. The vitamin mixture (per 100 mL) contains 100 mg of nicotinic acid, 100 mg of thiamine hydrochloride, 5 mg of biotin, 50 mg of p-aminobenzoic acid, 1 mg of vitamin B12, 50 mg of calcium pantothenate, 50 mg of pyridoxine hydrochloride, and 50 mg of folic acid. The trace mineral solution (per liter) contains 1.11 g of FeSO47H2O, 24.65 g of MgSO47H2O, 2.94 g of CaC122H2O, 23.4 g of NaCl, 111 mg of MnSO44H2O, 28.8 mg of ZnSO47H2O, 29.2 mg of Co(NO3)26H2O, 25.2 mg of CuSO45H2O, 24.2 mg of Na2MoO42H2O, 31.0 mg of H3BO4, and 4.53 g of trisodium EDTA. 5dPE medium [14] is a diluted form of PE medium containing fivefold reduced amounts (0.1 g/L) of organic compounds: sodium glutamate, disodium succinate hexahydrate, sodium acetate, yeast extract, and casamino acids. Anaerobic conditions are achieved by fully filling screw-capped glass tubes with the medium. 3. Solid medium: 1.5% (w/v) agar dissolved in PE or 5dPE medium. Anaerobic conditions are provided using an oxygen absorber. 4. Carbon-limited (CL) soft agar medium (per liter, pH 7.5): 0.05 g Na2S2O3, 0.38 g KH2PO4, 0.39 g K2HPO4, 0.05 g yeast extract, 1 mL of vitamin mixture (see above), 5 mL of trace mineral solution (see above), and 7 g of agar (0.7% w/v). 5. Heating incubator. 6. Illumination: tungsten incandescent lamp, 740 nm LED.

2.2

Microscopy

1. Tunnel glass slide: Coverslip (18  18  0.13–0.17 mm) is taped on a glass slide (25  75  1.0–1.2 mm) using two strips of double-sided tape (5 mm wide and over 18 mm long). Parallel and opposite sides of the coverslip are taped on the slide to create a void space in center (8  18 mm wide and

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Fig. 1 Cuvette holder equipped with an up-and-down motion device for adjusting the cuvette height. White arrows indicate the direction of beam through a 1 mm width slit

approximately 200 μm height). The glass slide can be replaced with a larger coverslip (24  60  0.13–0.17 mm) for efficient heating using a microscope stage heater (see below). 2. Glass beads: Micro-glass beads (1.32 μm of median size) are coated with 0.05% (w/v) poly-L-lysin solution. 3. Microscope with a digital camera: An upright phase-contrast microscope equipped with a 40  dry objective lens and a digital camera or a digital high-vision video camera. 4. Panel glass heater for microscope stage. 2.3

Spectroscopy

1. Photometer. 2. Cuvette holder: A handmade cell holder (Fig. 1) that can move a cuvette (external dimensions: 12  12  45–50 mm) up and down.

3

Methods For motility assays, cells in the exponential growth phase should be used (see Note 2).

3.1 Gliding Motility in Cell Suspension (CellAggregate Formation)

1. Suspend bacterial cells in 10 mM Tris–HCl, pH 8.0 (see Note 3). 2. Incubate the cell suspension at 55
C under illumination (see Note 4). 3. Observe the formation of cell aggregate (Fig. 2). Take record of the time taken for this process.

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Fig. 2 Cell-aggregate formation in C. aggregans NBF in a 1.5 mL plastic tube observed at various time intervals 3.2 Gliding Motility on Solid Medium

1. Suspend cells in a fresh medium at a final cell density of approximately OD660 ¼ 10. 2. Spot the dense cell suspension (10 μL, approximately 5 mm in diameter) on PE agar medium. 3. Incubate the agar plate under aerobic conditions at 55
C in the dark. 4. Dendritic formation is observed around the spot (Fig. 3). The expansion of the dendritic formation can be compared under different conditions. Chemo- and photo-tactic behaviors can be examined by the directional expansion (or suppression) of dendritic formation (see Note 5).

3.3 Gliding Motility in Solid Media

1. Inject 300 μL of carbon-limited (CL) soft agar medium into a glass cuvette (internal dimensions: 10  5  40 mm) and solidify it to prepare the first (bottom) layer. 2. Inject 400 μL of cell suspension with the CL soft agar medium over the first layer and solidify it to prepare the second (middle) layer. 3. Inject 300 μL of the CL soft agar medium over the second layer and solidify it to prepare the third (top) layer (Fig. 4a). A test chemical of interest (e.g., attractant and repellent) is added to the first or third layer. 4. Cap the cuvette with a rubber stopper and replace the gas phase by purging with N2 gas. 5. Incubate the cuvette at 55
C under illumination (see Note 4).

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Fig. 3 Colony morphology of C. aggregans NBF on PE agar medium

Fig. 4 Directional motility in agar medium in a spectrophotometer cuvette. (a) three-layered agar in a cuvette and (b) distribution of absorbance at 740 nm (ABS740) at each vertical position (representative examples). Time-series data (e.g., attractant in the bottom layer or repellent in the top layer) are shown as thick to dotted to dashed lines

6. Fix the cuvette in a handmade cuvette holder (see Subheading 2, Fig. 1), and measure the absorbance at 1 mm interval within and surrounding parts of the middle layer using a spectrophotometer. Absorbance at 740 nm is an index of the amount of photosynthetic pigment (bacteriochlorophyll c) of Chloroflexus. Effects of attractive and repellent chemicals in the top or bottom layer are detected by the changes in distribution of absorbance at 740 nm (Fig. 4b).

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Fig. 5 Sequential micrographs recorded at 1 min intervals. Arrowheads in the first photo indicate the moving direction of each filament. Scale bar, 50 μm 3.4 Gliding Motility on Microscopic Glass Slide

1. Inject cell suspension (approximately 20 μL) into a tunnel glass slide. 2. Incubate the glass slide at 55
C in light for a few minutes to allow cells to attach the glass surface (see Note 6). 3. Observe the cells using a phase-contrast microscope with a stage heater under illumination (Fig. 5) (see Note 7).

3.5 Cell-Surface Movements

1. Mix a culture solution with a suspension of glass beads. 2. Centrifuge the mixture (4000  g, 2 min) to remove the supernatant. 3. Suspend the precipitates gently in a fresh medium. 4. Inject the suspension (approximately 20 μL) into a tunnel glass slide. 5. Observe movements of glass beads on the cell surface using a phase-contrast microscope with a stage heater under illumination (Fig. 6). Beads on the cell-surface move along with the filaments at a speed comparable with the gliding speed, for example, 2–3 μm s1 for C. aggregans NBF at 55
C [11].

4

Notes 1. These cultures can be stored at 80
C in 10–15% (v/v) glycerol. 2. Exponential growth phase of Chlroflexus under anaerobic light conditions is generally measured to be an OD660 of approximately 0.3. Aerobic culturing in the dark is also useful.

Motility Assays of Chloroflexus

389

Fig. 6 Microscopic image showing a glass bead (approximately 1.2 μm diameter) (indicated by the white arrow) attached to a filamentous cell. Beads on the filaments of C. aggregans NBF moved 2–5 μm in one direction and occasionally reversed the direction [11]

Repetitive sub-culturing may reduce active motility. It is better to occasionally revive Chloroflexus cultures from frozen stock. 3. The OD660 should be 0.3–0.4. Remove air in a tube as much as possible (e.g., 1.4 mL of cell suspension in a 1.5 mL tube). Instead of plastic tubes, microtiter plates (96-well, 48-well, or 24-well) are also applicable. 4. The quality and quantity of illumination affect gliding motility of Chloroflexus. 30–50 W m2 by a tungsten lamp could be useful in the initial trials. LED illumination at 740 nm (e.g., 30 W m2) is also applicable. 5. The effects of chromatography resin on motility can also be examined. Interactions between the bacterial cell-surface and resins possibly affect the dendritic formation on the agar medium. Chromatography resin for ion-exchange and sizeexclusion is suspended in sterile water according to the manufacturer’s instructions and washed several times with sterile water. Prior to spotting the bacterial cells, the resin suspension is spread on the agar medium. As an example, the expansion of the dendritic formation of C. aggregans NBF is facilitated by DEAE-Sepharose (Cytiva, Tokyo, Japan). 6. Unattached cells on a glass slide may interfere with the precise microscopic observation. Unattached cells are washed away from the tunnel slide by flowing an appropriate amount of fresh medium prior to observation. For long-term observations (e.g., longer than 5 min), a cover slip should be sealed using clear nail polish. 7. The reversal frequency of each filament is counted during a long-term observation, (e.g., 10–20 min).

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Acknowledgments We thank Satoshi Hanada and Daisuke Nakane for their valuable suggestions. This work was partially supported by JSPS KAKENHI Grant Number JP21K06338 to S.H. References 1. Pierson BK, Castenholz RW (2015) Chloroflexus. In: Trujillo ME, Dedysh S, DeVos P, Hedlund B, Ka¨mpfer P, Rainey FA, Whitman WB (eds) Bergey’s manual of systematics of archaea and bacteria. Wiley, New York 2. Oren A, Garrity GM (2021) Valid publication of the names of forty-two phyla of prokaryotes. Int J Syst Evol Microbiol 71:005056 3. Blankenship RE (2001) Molecular evidence for the evolution of photosynthesis. Trends Plant Sci 6:4–6 4. Kawai S, Nishihara A, Matsuura K et al (2019) Hydrogen-dependent autotrophic growth in phototrophic and chemolithotrophic cultures of thermophilic bacteria, Chloroflexus aggregans and Chloroflexus aurantiacus, isolated from Nakabusa hot springs. FEMS Microbiol Lett 366:fnz122 5. Kanno N, Haruta S, Hanada S (2019) Sulfidedependent photoautotrophy in a filamentous anoxygenic phototrophic bacterium, Chloroflexus aggregans. Microbes Environ 34:304– 309 6. Hanada S, Hiraishi A, Shimada K et al (1995) Isolation of Chloroflexus aurantiacus and related thermophilic phototrophic bacteria from Japanese hot springs using improved isolation procedure. J Gen Appl Micriobiol 41: 119–130 7. Hanada S (2014) The phylum Chloroflexi, the family Chloroflexaceae, and the related phototrophic families Oscillochloridaceae and Roseiflexaceae. In: Rosenberg E, Delong EF, Lory S et al (eds) The prokaryotes, vol 11, 4th edn. Springer, New York

8. Miyata M, Robinson RC, Uyeda TQP (2020) Tree of motility – a proposed history of motility systems in the tree of life. Genes Cells 25:6–21 9. Hanada S, Hiraishi A, Shimada K et al (1995) Chloroflexus aggregans sp. nov., a filamentous phototrophic bacterium which forms dense cell aggregates by active gliding movement. Int J Syst Bacteriol 45:676–681 10. Hanada S, Shimada K, Matsuura K (2002) Active and energy-dependent rapid formation of cell aggregates in the thermophilic photosynthetic bacterium Chloroflexus aggregans. FEMS Microbiol Lett 2208:275–279 11. Fukushima S, Morohoshi S, Hanada S et al (2016) Gliding motility driven by individual cell-surface movements in a multicellular filamentous bacterium Chloroflexus aggregans. FEMS Microbiol Lett 363:fnw056 12. Klatt CG, Bryant DA, Ward DM (2007) Comparative genomics provides evidence for the 3-hydroxypropionate autotrophic pathway in filamentous anoxygenic phototrophic bacteria and in hot spring microbial mats. Environ Microbiol 9:2067–2078 13. Morohoshi S, Matsuura K, Haruta S (2015) Secreted protease mediates interspecies interaction and promotes cell aggregation of the photosynthetic bacterium Chloroflexus aggregans. FEMS Microbiol Lett 362:1–5 14. Hirose S, Matsuura K, Haruta S (2016) Phylogenetically diverse aerobic anoxygenic phototrophic bacteria isolated from epilithic biofilms in Tama River, Japan. Microbes Environ 31: 299–306

INDEX A Achromaticity ................................................................ 221 Acquisition.........................................................37, 88, 89, 222–224, 227, 239, 242, 365 Adenosine triphosphate (ATP).................... 4, 19, 20, 23, 27, 183, 197, 200, 206, 207, 311, 328–330, 333, 334, 360, 363, 369, 370 Adherence........................................................................ 41 Adhesin SprB ....................................................... 268, 275, 278 Aerotaxis ........................................................................ 134 Agar Bacto ..................................... 6, 19, 45, 148, 288, 289 Eiken ...................................................... 148, 150, 155 Agarose gel electrophoresis ................................. 351, 354 Amino acid cysteine ................................. 30, 73, 75, 80, 105, 207 isotope-labeled phenylalanine .................................. 60 Amino acid precursor α-ketobutyric acid ........................................ 60, 62, 66 Aminoacyl-tRNA synthase........................................72–74 2-amino-6-mercapto-7-methylpurine riboside (MESG)........................................................ 360 Amphipols A8-35................................................. 112, 114 Angstrom (Å) ............................................................49, 51 Antibody anti-FlgD ................................................................... 24 anti-FlhA.................................................................... 24 anti-FliG .................................................................... 73 anti-FliO ...................................................................... 8 anti-FliP ....................................................................... 8 anti-His-tag ............................................................. 114 anti-lipopolysaccharide (LPS) antiserum ............... 166 anti-MotP ................................................................ 122 anti-MotS................................................................. 122 anti-PomA ................................................................. 73 antisera against SprB ............................................... 280 fluorescently labeled anti-rabbit IgG ..................... 280 horseradish peroxidase (HRP) conjugated anti-rabbit IgG antibody (Goat anti-mouse IgG-HRP) .................................. 77 monoclonal anti-FLAG antibody............................. 13 monoclonal anti-HA antibody .............................8, 13

Antifungal toxins........................................................... 250 Aperture...................................... 219–221, 237, 239, 261 Archaea Halobacterium salinarum.....................192, 199–202, 206, 207 Haloferax volcanii ......................... 193, 199–202, 206 Methanocaldococcus villosus ..................................... 184 Methanococcus M. jannaschii .................................................72, 79 Methanoculleus................................................ 185, 191 Methanogenium .............................................. 185, 191 Methanospirillum hungatei ..................................... 191 Methanothermococcus M. thermolithotrophicus ..................................... 192 Natrialba magadii.................................................. 191 Pyrococcus furiosus ................................................... 191 Saccharolobus shibatae ............................................. 191 Sulfolobus acidocaldarius............................... 184–188, 191–193 Thermoplasma volcanium ....................................... 191 Archaeal adhesion pili (Aap pili) .................................. 191 Archaellar protein archaellin............................... 183, 184, 189, 192, 206 ArlA (FlgA1)............................................................ 197 ArlB .......................................................................... 197 ArlC/D/E............................................................... 197 ArlF .......................................................................... 198 ArlG ......................................................................... 198 ArlH ......................................................................... 198 ArlI ........................................................................... 199 ArlJ .................................................................. 197, 199 Archaellum (archaella) ............................... 183–192, 197, 199, 204, 206 Assembly flagellar....................................................................... 36 stator ....................................................................72, 96 Atomic model............................................................51, 52 ATPase FoF1-ATP synthase F-type ATPase, α-subunit ................................. 328 F-type ATPase, β-subunit ................................. 328 Attachment organelle........................................... 352, 356 Attenuated total reflectance-Fourier transform infrared (ATR-FTIR) spectroscopy ....................95–106

Tohru Minamino et al. (eds.), Bacterial and Archaeal Motility, Methods in Molecular Biology, vol. 2646, https://doi.org/10.1007/978-1-0716-3060-0, © The Editor(s) (if applicable) and The Author(s), under exclusive license to Springer Science+Business Media, LLC, part of Springer Nature 2023

391

BACTERIAL AND ARCHAEAL MOTILITY

392 Index

Autofluorescence..................................... 89, 92, 217, 218 AutoGrid C-clip ....................................................................... 216 C-clip ring ...................................................... 215, 216 holey carbon film grid............................................. 213 AutoGrid assembly workstation .......................... 213, 215 AutoGrid forceps.................................................. 216, 217 Autoloader................................................... 213, 215, 217 Autoloader cassette ....................................................... 217 Avidin.................................................................... 338, 341

B Back-projecting ............................................................. 212 Bacteria anoxygenic photosynthetic bacteria ....................... 383 Bacillus B. clausii .............................................................. 84 B. subtilis...........................................110, 120, 148 Bacteroidetes (Bacteroidota) Flavobacterium columnare ................................ 287 Flavobacterium johnsoniae .......................268–275, 277, 287 Flavobacterium psychrophilum .......................... 287 Porphyromonas gingivalis .................................. 269 Saprospira grandis str. Lewin .................. 269, 274 Borrelia burgdorferi................................................. 159 Brachyspira hyodysenteriae ....................................... 159 Campylobacter jejuni................................................. 84 Chloroflexus aggregans .................................. 383, 384, 386, 388, 389 cyanobacteria .................................................. 255, 256 Escherichia coli auxotrophic E. coli ........................................66, 67 E. coli BL21 (DE3) ......................................60, 66, 86, 92, 102, 105, 111 E. coli DB3.1 ..................................................... 349 E. coli DH5α ...........................300, 348, 349, 351 E. coli S17-1 .....................................134, 137, 138 E. coli WM3064 ...............................134, 137, 138 filamentous bacteria .................................17, 360, 384 gram negative .......................................................... 287 gram positive ........................................................... 321 Lysobacter enzymogenes ............................................ 250 Magnetospirillum magneticum AMB-1 ................. 134 Mollicutes hominis, Mycoplasma mobile ............................. 321 phytoplasma....................................................... 321 pneuomiae, pneumoniae ................................... 321 Spiroplasma, Spiroplasma eriocheiris ............... 374, 375, 377–379 myxobacteria Myxococcus xanthus..................211, 213, 214, 217 Proteus mirabilis ...................................................... 148 Pseudomonas aeruginosa ................................ 148, 249

Salmonella enterica serovar Typhimurium ........3, 127 Spirochete Leptospira biflexa ............................................... 173 Leptospira interrogans ....................................... 159 Leptospira kobayashii ......................................... 173 Synechocystis sp. PCC6803 .......................... 256–262 Thermosynechococcus vulcanus ................256, 258–261 Vibrio V. alginolyticus....................................... 36, 38, 39, 74, 75, 79, 80, 86, 96 V. parahaemolyticus ........................................... 148 wall-less .................................................................... 321 xanthomonadaceae.................................................. 250 Bacterial flagellum (bacterial flagella) axial structure ............................................... 17, 18, 96 basal body C-ring ............................................................36, 40 L-ring .............................................................36, 96 MS-ring............................................................ 4, 17 P-ring .............................................................36, 96 rod..................................................................18, 96 filament L-type straight filament ...................................... 44 R-type straight filament ...................................... 44 sticky filament........................................... 126, 130 hook .....................................................................18, 35 hook-basal body (HBB) ........................................... 84 periplasmic flagellum............................................... 160 peritrichous flagellum ............................................. 147 polar flagellum........................................................... 36 Bacterial nanomachines................................125, 211–244 Bacterial organelle ................................................ 133, 134 Basal body................................................. 3, 4, 17, 25, 36, 43, 83, 211, 212, 267 Bead assay carboxylated polystyrene bead gold nanoparticle .............................................. 130 fluorescent bead latex bead ........................................................... 174 microsphere (bead) ........................................... 258 streptavidin-conjugated bead carboxylated-polystyrene bead ............... 199, 202, 338, 341 1-(3-dimethylaminopropyl)-3-ethylcarbodiimide hydrochloride (EDC).................................. 338 N-hydroxysulfosuccinimide sodium salt (Sulfo-NHS) ................................................ 200 Beam beam induced movement beam shift ................................................. 219, 220 beam tilt............................................................. 219 B-factor ............................................................................ 51 Binning ................................................... 49–51, 222–224, 228, 231, 233

BACTERIAL Bio-bead .....................................102, 104, 112, 115, 123 Biofilm ........................................211, 249, 287, 288, 296 Biotin biotin-NHS ester............................................ 199, 202 biotin-PEG2-maleimide ................................ 199, 202 sulfo-NHS-LC-LC-biotin ...................................... 340 Biotinylated cell............................................................. 202 Blotting...................................................8, 13, 49, 76, 78, 114, 120, 122, 214, 215, 239, 271–273, 354 Boltzmann constant ............................................. 172, 342 Border-crossing assay ........................................... 149, 156 Bovine serum albumin (BSA)...........................7, 11, 122, 162, 173, 199, 203, 207, 214, 239, 329, 330, 339, 341, 344, 375–377 Bromophenol blue ......................... 8, 21, 45, 75, 76, 314 Brownian motion ........................................ 268, 342, 343 Bundle.............................................................44, 274, 275 Bundled filaments ......................................................... 268

C Camera charged coupled device (CCD) camera ............46, 48, 164, 200, 280, 281, 283, 304, 314, 324, 330 complementary metal oxide semiconductor (CMOS) camera ................................201, 258, 330, 376 digital camera .......................................................... 385 electron multiplying CCD (EMCCD) camera....... 39, 136, 141, 200, 203, 289, 328, 330, 331 FEI Falcon II 4k × 4k CMOS direct electron detector camera ............................................................ 46 Gatan K3 Summit camera ............................. 213, 214 high-speed camera................................................... 128 scientific CMOS (sCMOS) camera ...................39, 90, 204, 330, 332 TVIPS TemCam-F415MP 4k × 4k CCD camera... 46 video camera................ 128, 130, 163, 174, 177, 385 Carbon ............................................. 46, 48, 57, 186, 192, 213–217, 219, 220, 226, 239, 290, 294, 295, 303, 305 Carbon-grid stamp-peel method................................... 48, 289–290, 294–295 Carbonyl cyanide 3-chlorophenylhydrazone (CCCP) ...........................87, 90–92, 280, 281, 284, 375, 376, 378 Cartridge .............................................................. 216–218 CBB staining ........................................... 11, 12, 116, 316 Cell aggregation ................................................................ 51 density................................................. 9, 10, 156, 204, 206, 251, 287, 314, 386 elongation................................................................ 148 morphology .................................................... 299, 374 wall ............................................................95, 299, 359

AND

ARCHAEAL MOTILITY Index 393

Centrifuge.......................................6, 7, 9–11, 19–21, 39, 46, 51, 61, 75–78, 88–90, 103, 111–114, 129, 138, 140, 141, 144, 163, 164, 174, 175, 185, 186, 189, 191, 201, 202, 207, 214, 251, 269, 289, 292–294, 303, 313–316, 322, 331, 341, 349, 350, 354, 355, 361–364, 366, 388 Chemiluminescence ................................ 8, 13, 21, 77, 79 Chemotaxis chemotaxis plate ...................................................... 156 chemotactic signaling................................................ 57 chemotactic signaling protein CheY .................................................................... 57 Chimeric protein PotB .................................................74, 75, 77, 79, 80 Chromatography Co-TALON affinity chromatography ...................... 98 fast protein liquid chromatography (FPLC) system...................................... 7, 99, 104, 112, 313, 362, 364, 369 Ni-NTA affinity chromatography............................. 91 size exclusion chromatography (SEC) (gel filtration) ............................... 7, 12, 61, 62, 64, 67, 96, 99, 112, 115, 121 Classification three-dimensional (3D) classification ................50, 51 two-dimensional (2D) classification................ 49, 312 Clustering K-means ...............................................................30, 32 CO2 incubator............................................................... 163 Cold shock................................................... 51, 66, 96, 99 Collodion.............................................256, 258, 260–262 Colonization........................................147, 157, 249, 250 Colony non spreading colony..................................... 287, 297 spreading colony ................................... 287, 295, 296 Conductivity .................................................................. 250 Conjugation (conjugative transfer).................... 134, 137, 141, 212, 289, 297 Contrast transfer function (CTF) ...........................49–51, 233, 317 Coomassie Brilliant blue (CBB) staining........................ 8, 122, 315 Cross-linking disulfide cross-linking oxidant, Copper (II) phenanthroline................. 73 photo-crosslinking amber stop codon ............................................... 73 amber suppressor tRNA ..................................... 72 p-benzoyl-L-phenylalanine ( pBpA) .............72, 73 Cryogen container ........................................................ 215 Cryo-grid box....................................................... 215, 216 Cryo tools dryer ............................................................ 213 Cryo transfer container ................................................. 215 Crystallization ............................................................... 275

BACTERIAL AND ARCHAEAL MOTILITY

394 Index

CTF refinement.........................................................50, 51 Culture density.............................................................. 154 Culture media Aluotto medium............................................ 313, 314, 322, 323, 329, 331, 339, 340 antibiotics amphotericin B ......................................... 313, 322 ampicillin .............................................6, 9, 19, 74, 76–79, 86, 88, 99, 102, 111, 288, 292, 293, 313, 322, 329, 339, 348, 349, 351, 361, 363 chloramphenicol..............................19, 74, 77, 79, 86, 88, 349 erythromycin ............................................ 289, 297 gentamicin ...................... 349, 350, 352, 353, 355 kanamycin ....................................... 135, 136, 138, 139, 152, 348, 349 penicillin G ........................................................ 339 streptomycin ...................................................... 289 arabinose (L-arabinose) .............................. 19, 21, 74, 76–78, 80, 86 Balch medium Balch medium III....................185, 189, 191, 192 BG11 medium...............................257, 258, 260, 261 Brock’s medium Brock I ...................................................... 184, 187 Brock II/III ............................................. 184, 187 carbon-limited soft agar medium.................. 384, 386 casamino acid solution (Ca-solution) ........... 155, 200 Casitone-yeast extract (CYE) agar plate CYE-Ca agar plate............................289, 292, 293 CYE-Em agar plate ...................................292–294 CYE-Sm agar plate ............................................ 293 Casitone-yeast extract (CYE) medium .................. 288 Eagle’s minimum essential medium (MEM)........ 163, 164, 166 Ellinghausen–McCullough–Johnson–Harris (EMJH) liquid medium ................... 162–164, 173, 175 freeze throw buffer (FTB) ............................. 289, 292 heart infusion broth ...................................... 313, 322, 324, 329, 339, 375 heat-inactivated horse serum ......................... 313, 339 horse serum .......................................... 322, 324, 329, 339–341, 344 isopropyl-β-D-thiogalactopyranoside (IPTG) .................................................. 86, 111 LB agar (LA) plate (Luria-Bertani agar).............6, 19, 21, 45, 251, 292, 293, 348 LBG medium.................................................. 111, 113 L-broth (Lennox broth, Luria broth, Luria-Bertani broth, Lysogeny broth, Lysis broth) (LB)...............................................86, 111, 127 M9 medium.........................................................60, 66 Magnetospirillum growth medium (MSGM) ....... 135, 136, 138–141

marine broth..................................269, 271, 273, 275 motility buffer (Motility medium, MM) ..........86–91, 200–204, 279, 376 PE medium..................................................... 384, 388 peptone yeast 2 agar plate (PY2) .......................... 289, 294, 297 peptone yeast glucose (PYG) ........................ 289, 294 PPLO agar plate .................................... 352, 353, 355 PPLO broth.......................... 350, 352, 353, 355, 356 R2 medium.............................................................. 377 soft agar ................................................ 148, 149, 151, 152, 154–156, 288 SP-4 medium.................................................. 339, 340 swarm agar............................................. 150, 152, 153 T-broth (TB) ......................................... 86, 88, 91, 92 TC buffer ................................................289, 292–294 TG broth ...................................................... 74, 76–80 TMK buffer .........................................................37, 39 TMN buffer.........................................................37, 39 tryptic soy broth............................................. 250, 251 tryptic soy broth agar.............................................. 251 VC medium ......................................................... 37–39 VPG medium ............................................................ 37 2YT ........................................................................6, 10 Cultured cells MDCK-NBL2 (dog kidney epithelial cell)........... 163, 166 NRK-52E (rat kidney epithelial cell) ..................... 166 Cytoplasmic membrane ..................................3, 4, 17, 18, 83, 126, 279 Cytoskeletal protein actin superfamily fibril........................................................... 359, 373 HMW2 .............................................................. 352 MreB .................................................359–370, 373 Cytoskeleton (cytoskeletal structures) ............... 141, 321, 328, 369

D Defocus.................................................49, 219, 220, 222, 223, 225–227, 229, 231, 242 Density map............................................................ 51, 318 Desalting column .......................................................... 199 Desiccator .......................................................8, 13, 46, 48 Detergent 3-[(3-Cholamidopropyl)-dimethylammonio]-1Propane Sulfonate] (CHAPS) ...................... 98 decyl maltose neopentyl glycol (DMNG) .............. 96, 103, 115, 121, 123 decyl maltoside (DM) .................................... 115, 121 lauryl maltose neopentyl glycol (LMNG) ............... 96 n-dodecyl-β-D-maltoside (Dodecyl maltoside, DDM) .......................................................... 270 OP-10 ............................................................. 186, 189

BACTERIAL sodium dodecyl sulfate (SDS) .................................. 96 triton X-100 ......................................... 20, 24, 25, 45, 47, 269, 272, 312, 313, 315, 316, 318, 329, 330, 335 Deuterium oxide (D2O) ...........................................60, 61 Dextrin.................................................................. 185, 187 Diaminopimelate (DAP)............................. 135, 138, 144 Diderm........................................................................... 300 1-(3-dimethylaminopropyl)-3-ethylcarbodiimide (EDAC)........................................................ 177 Dithiothreitol (DTT)......................................20, 23, 329, 330, 363, 366, 370 DNA ligation........................................................ 349, 354 Domains ........................................ 19, 27, 29, 31, 44, 58, 85, 96, 117, 123, 130, 197 Dose fractionation....................................... 223, 224, 228 Drag coefficient ...................................127, 169, 207, 332 Drift ........................................................................ 49, 105 Dynamics .............................................. 28, 32, 35, 57–59, 84, 110, 111, 118, 121, 126, 133, 134, 166, 211, 255–257, 262, 278, 360, 373

E Electron crystallography ............................................... 212 Electron dose..................................................49, 228, 229 Energy-conversion efficiency ........................................ 329 Energy filter ................................................. 220, 223, 230 Enzyme DNA polymerase ................................... 289, 349, 356 DNase ................................................... 186, 190, 200, 313, 329, 330 lysozyme ................................. 18, 19, 22, 45, 47, 105 LR clonase ...................................................... 349, 350 protease chymotrypsin ............................................ 300, 303 proteinase K........................................19, 349, 350 restriction enzyme BamHI ..............................................289, 292, 293 HindIII ..................................................... 349, 354 NotI .......................................................... 289, 293 RNase........................................................ 186, 189 SalI ............................................................ 289, 292 SphI.................................................................... 289 T4 DNA ligase .................................................. 354 Epon812 ............................................................... 291, 296 Ethane/propane................................................... 213, 215 Ethylenediaminetetraacetic acid (EDTA) Ethylenediamine-N, N, N’, N’-tetraacetic acid, dipotassium salt, dihydrate (EDTA 2K) ...... 87 1-Ethyl-3-(3-Dimethylaminopropyl) carbodiimide (EDAC)............................................... 177, 338 Eucentric focus.............................................................. 219 Eucentric height............................................................ 219 Eucentricity ................................ 219, 225, 227–230, 243

AND

ARCHAEAL MOTILITY Index 395

Eukaryote ............................................................. 197, 329 Euler angle...........................................235, 236, 238, 240 Euryarchaeota................................................................ 197 Exposure time ............................................ 37, 39, 49, 90, 140, 149, 150, 218, 222–224, 228, 281, 282, 316, 377, 379

F Fiducial .......................................214, 215, 232, 233, 239 Filamentous fungi ......................................................... 250 Flagellar ejection ................................................ 36, 39–41 Flagellar motor proton (H+)-driven motor................ 71, 96, 268, 279 rotary motor .......................................... 43, 71, 83, 95 rotor ........................................ 71, 83, 84, 95, 96, 110 sodium ion (Na+)-driven motor.........................71, 96 stator rotor interaction .......................................72, 96 stator .................................58, 71, 83, 84, 95–97, 110 Flagellar motor rotation clockwise (CW) ......................................................... 57 counterclockwise (CCW).......................................... 57 Flagellar outer membrane complex (FOMC) ............... 36 Flagellar proteins flagellin FliC .............................................. 50, 51, 127, 130 FljB....................................................47, 48, 50, 51 rod-hook type protein ........................................ 25 FlgD ........................................................19–21, 23, 25 FlgE.......................................................................... 127 FlgG ......................................................................... 127 FlhA ..................................4, 5, 17, 21, 23, 25, 27, 28 FlhB ...............................................4, 5, 17, 19, 21, 25 FlhC ........................................................................... 25 FlhD........................................................................... 25 FliF .........................................................................4, 72 FliG ................................................. 40, 57, 58, 72–75, 77, 78, 80, 85 FliH........................................................................4, 17 FliI............................................................4, 17, 20, 23, 25, 250, 252, 253 FliJ...................................................... 4, 17, 20, 23, 25 FliK .......................................................................... 127 FliM .............................................................. 57, 58, 72 FliN ................................................................. 4, 57, 72 FliO ............................................................. 5, 8, 11–14 FliP ...............................................4, 5, 7, 8, 11–14, 17 FliQ ..............................................4, 5, 7, 8, 11–14, 17 FliR ..............................................4, 5, 7, 8, 11–14, 17 FliT................................................................ 19, 21, 25 MotA........................................ 71, 75, 79, 80, 83–86, 89, 90, 92, 96, 110, 279 MotB............................................... 71, 79, 83–86, 90, 92, 96, 110, 279 MotP ......................................................... 84, 109–123

BACTERIAL AND ARCHAEAL MOTILITY

396 Index

Flagellar proteins (cont.) MotS .............................. 84, 111, 115, 117, 120, 122 PomA .....................................................71, 73–80, 84, 86, 91, 96, 99, 103, 105, 106 PomB ..............................................71, 79, 84, 86, 91, 96, 97, 99, 103, 105, 106 Fluorescence spectrophotometer ........................... 86, 88, 113, 118, 123 Fluorescent dye CoroNa Green-AM.....................85, 89–92, 113, 118 Cy3.................................................................. 207, 331 Dylight 488 ............................................................. 199 FM® 4-64 dye (N-(3-Triethylammoniumpropyl)-4(6-(4-(Diethylamino) Phenyl) Hexatrienyl) Pyridinium Dibromide) .............................. 376 streptavidin-conjugated fluorescent dye ....... 199, 202 Fluorescent protein eGFP-FliG ...........................................................37, 38 enhanced green fluorescent protein (eGFP) ........... 40 enhanced yellow fluorescent protein (EYFP) ........ 352 green fluorescent protein (GFP) .......... 166, 288, 293 pH-sensitive fluorescent protein E2GFP...............................................140, 143, 144 pHluorin(M153R) .......................................85, 86, 88–90, 92, 93 Force field ff14SB ........................................................................ 30 ff19SB ........................................................................ 30 ff99SBildn.................................................................. 30 SPC/Eb water ........................................................... 30 Fourier pixel ......................................................... 239, 244 Fourier Shell Correlation (FSC) ...................51, 236, 239 Frame ....................................................49, 117, 119, 120, 129, 130, 164, 166, 177, 204, 206, 207, 228–231, 243, 283, 324, 332, 334, 338, 342 Free energy .................................28, 29, 31, 32, 161, 197 Freeze-etching device JFDV............................................................... 302, 305 Freezing machine CryoPress................................................................. 301 Fresh yeast extract ....................................... 339, 348, 354

G Gain reference ...................................................... 221, 242 Gas vacuoles ......................................................... 201, 207 Gateway cloning...........................................348–351, 354 Gaussian function................................117, 175, 332, 334 Gear rotation ................................................................... 72 Gellan gum .................................................. 136, 139, 144 Genetic manipulation.................................................... 347 Ghost gliding ............................................................. 331, 334 rotary ....................................................................... 332 Gliding machinery..............................267–269, 275, 288, 312, 383, 384

Gliding protein gldA ......................................................................... 287 gldB .......................................................................... 287 gldD ......................................................................... 287 gldF .......................................................................... 287 gldG.......................................................................... 287 gldH ......................................................................... 287 gldI........................................................................... 287 gldJ ........................................................................... 287 gldK ......................................................................... 287 gldL .......................................................................... 287 gldM ......................................................................... 287 gldN ......................................................................... 287 gldO ......................................................................... 287 Gli123 ............................................................. 327, 328 Gli349 .................................................... 311, 321, 327 Gli521 ............................................................. 327, 328 Glow-discharge glow discharging cleaning system .......................... 213 Glucose ........................................37, 111, 148, 150, 152, 153, 155, 288, 289, 297, 330, 339–343, 348 Glutaraldehyde ........................................... 193, 270, 271, 275, 290, 291, 295, 296 Glycan chain .................................................................. 299 Glycosylation N-glycosylation ....................................................... 183 O-glycosylation........................................................ 183 Gold colloid.......................................................... 213, 214 Gold standard......................................................... 51, 236 Gradient fractionator ................................................46–48 Gradient master.........................................................46, 47 Gramicidin .......................................................... 87, 90–92 Grating spectrometer .................................. 136, 141, 142 Grid screening ...................................................... 216–218

H Hair-like protein appendage......................................... 249 Hard agar....................................................................... 148 Helical symmetry axial rise ..................................................................... 44 twist angel.................................................................. 44 Helical track................................268, 275, 277, 278, 283 Helix body helicity .......................................... 374, 375, 379 left-handed helix (LH)................................... 373, 378 right-handed helix (RH)....................... 283, 373, 378 High-pressure homogenizer French Press ....................................98, 102, 103, 105 Hole ..................................................................... 219, 221, 225, 227, 228, 230 Horizontal gene transfer............................................... 249 Humidity chamber ...................................... 156, 322, 323 Hydration ............................................................. 148, 155

BACTERIAL I Ice thickness ........................................216, 218, 225–227 Illumination laser .......................................................................... 198 LED ..........................................................37, 259, 389 mercury light source system ..................................... 87 tungsten lamp.......................................................... 389 Image distortion ........................................................... 221 Image shift..................................................................... 227 Immune system ...................................................... 44, 299 Immunoblot (western blot) ........................ 8, 12, 13, 19, 21, 23, 24, 72–78, 92, 105 Immunofluorescent labeling ............................... 278, 279 Infection .........................................................43, 147, 171 Initial model .....................................................50, 51, 318 In situ structure determination ........................... 211–244 Intensity ................................................37, 39, 68, 80, 85, 89–92, 115, 118, 130, 142, 143, 204, 205, 220, 221, 223, 255, 257, 334 Intracellular pH............................................................... 85 Inverted membrane vesicle (IMV) ..............18–20, 22–25 Inverted repeat (IR)...................................................... 355 Ion coater ....................................................................8, 46 Ion influx ......................................................................... 97 Ion motive force.............................................................. 95 Isothermal-isobaric (NPT) condition ............................ 30

K Kink......................................................373, 374, 377, 379 Kymograph ................................................. 141, 176, 203, 204, 256, 261, 278, 281, 283

L Lipopolysaccharide (LPS) ............................................. 166 Lipoprotein........................................................... 268, 277 Liquid ethane .................................................................. 49 Liquid nitrogen .......................................... 20, 23, 49, 96, 215, 216, 300–302, 304, 307 Low-pass filter ................................................51, 120, 232 Lowry method......................................... 10–11, 114, 122 LR reaction .................................................................... 351

M Macromolecular complex ............................................. 212 Magnetosome....................................................... 133–144 Magnetosome membrane protein MamC ...................................................................... 137 MamI ....................................................................... 137 Mms6 .............................................................. 140, 144 MmsF....................................................................... 137 Magnetotactic bacteria.................................................. 134 Magnetotaxis ........................................................ 133–144

AND

ARCHAEAL MOTILITY Index 397

Magnification ....................................... 49, 217, 221–228, 230, 233, 239, 242, 252, 294, 316, 352 Markov state model (MSM) ....................... 28, 29, 31, 32 Mean-square displacement (MSD) ..................... 161, 162 Membrane ...................................................3–6, 8, 10–13, 17, 18, 21, 23, 25, 36, 43, 47, 72, 76, 78, 79, 83, 84, 95–97, 99, 103, 111, 114, 115, 122, 126, 133–135, 137, 138, 147, 189, 228, 233, 234, 267, 275, 278, 279, 299, 300, 311, 312, 318, 321, 334, 335, 340, 349, 352, 353, 362, 364, 365, 378 Membrane proteins...........................................27, 83, 96, 114, 120–122, 137–141, 144 Methylcellulose ...................................329, 330, 332, 376 Mica flake.............................................................. 301, 304 Micro-optics effect ........................................................ 261 Microscope bright-field microscopy........................................... 261 carbon-coated copper 400 mesh grids................... 314 cover slip ......................................................... 322, 323 cryo-light microscope fluorescence channel .................................216–218 transmitted light brightfield channel (TL-BF) ....................................................... 217 dark field microscopy ..................................... 166, 177 electron cryomicroscopy (cryo-electron microscopy, cryoEM) cryo-transmission microscope (cryo-TEM) ....................... 212, 213, 215–217 plunge-freezing device ........................................ 46 single particle image analysis ........................44, 46 electron cryotomography (cryo-electron tomography, cryo-ET) subtomogram averaging .......................... 212, 213 electron microscopy (EM) atmospheric scanning electron microscopy (ASEM) dishes...................290, 291, 295, 296 quick-freeze-replica electron microscopy ........ 268 EM grid ................................................................... 212 epi-illuminator......................................................... 378 fluorescence ........................................... 280, 283, 294 fluorescence channel transmitted light brightfield channel (TL-BF) ....................................................... 217 fluorescence microscopy epifluorescence microscopy .............................. 280 highly inclined and laminated optical sheet (HILO) microscopy ........................... 139, 140 immunofluorescence microscopy ..................... 283 time-lapse..........................................139, 140, 294 total internal reflection fluorescence (TIRF) microscopy.................................136, 140–142, 203, 283 glass chamber .......................................................... 259

BACTERIAL AND ARCHAEAL MOTILITY

398 Index

Microscope (cont.) glass slide ........................................... 8, 167, 388, 389 GoldEnhance-EM thiocarbohydrazide (TCH) .............................. 290 high-speed atomic force microscopy (HS-AFM) cantilever..................................113, 115, 118, 121 mica disk ..................................113, 115, 117, 118 syringe pump system................................ 113, 121 inverted microscope ................................38, 136, 140, 200, 258, 330, 376, 379 light microscopy ............................................. 249, 321 negative stained EM phosphotungstic acid (PTA) ......................48, 303 uranyl acetate............................................ 191, 316 optical filter....................................258, 280, 282, 376 optical microscopy ........................................ 250, 256, 258, 280, 282, 288, 289, 291, 337, 376 phase-contrast microscopy............................ 201, 280, 312, 322, 324, 328, 330, 332, 334, 352, 374–377, 385, 388 stereomicroscope..................................................... 353 thermoplate ............................................................. 258 transmission electron microscope energy-filtered transmission electron microscopy (EFTEM) ............................................ 220, 239 tunnel slide (tunnel chamber) ........................ 38, 323, 324, 342 Microtome ..................................................................... 291 Minimum dose system .................................................... 49 Monomer....................................27, 29, 51, 96, 360, 365 Montage map ......................................223, 225–228, 242 Motility adhesion-based motility .......................................... 268 crawling motility ..................................................... 160 gliding motility..................................... 268, 269, 274, 277–279, 282–284, 287, 299, 311, 321, 327–335, 337, 344, 383, 389 social motility ................................................. 287–297 swimming motility ........................................ 149, 151, 152, 197, 373, 375 twitching motility..........................249, 250, 252, 255 Motor......................................................3, 13, 35, 36, 43, 44, 57–59, 71, 72, 83–85, 95–97, 109, 110, 118, 120, 125–130, 147, 148, 160, 183, 197–199, 204, 206, 207, 212, 268, 269, 279, 312, 315, 317, 318, 327–329, 334, 337 Motor chain..................................................312, 315–317 Motor protein kinesin ............................................................. 197, 334 myosin.................................................... 111, 197, 334 Motor speed .................................................................. 130 Motor switching...................................... 44, 58, 147, 198 Mutagenesis site-directed mutagenesis.......................................... 64 targeted mutagenesis ................................................ 73

N N-acetylneuraminyllactose (sialyllactose)..................... 329 NanoCab ....................................................................... 217 N-Butyl glycidyl ether (QY-1) ..................................... 291 NMR experiments HMQC ................................................................63, 67 NOESY-HMQC ....................................................... 66 NOESY-TROSY........................................................ 66 SOFAST-HMQC ...................................................... 67 TROSY ...................................................................... 65 NMR sample tube.....................................................61, 63 NOE ................................................................... 63, 65–68 Nuclear magnetic resonance (NMR) spectroscopy.............................................57–68 Numerical aperture (NA) .................................... 177, 338 NZ-amine ............................................................. 185, 187

O Objective......................................37, 128, 136, 152, 153, 155, 163, 167, 174, 177, 198, 200, 201, 207, 216, 217, 219, 220, 239, 252, 258–260, 280, 282, 322, 324, 330, 337, 338, 376, 379, 385 Offset ............................................................................. 226 Optical tweezers high numerical aperture objective lens .................. 338 lens heater................................................................ 338 mirror....................................................................... 338 Nd:YAG laser........................................................... 338 neutral density (ND) filter...................................... 338 piezoelectric stage ................................................... 338 plano convex lens .................................................... 338 stage heater.............................................................. 338 Osmium tetroxide (OsO4) ......................... 290, 291, 295 Osmotically shocked cell...................................... 273–275 Outer membrane........................................ 18, 36, 47, 51, 96, 126, 160, 166, 233, 234, 268, 274, 277–279, 299, 300

P 1-palmitoyl-2-oleoyl-sn-phophatidylethanolamine (POPE) ............................................... 102, 103 1-palmitoyl-2-oleoyl-sn-phosphatidylglycerol (POPG) ............................................... 102, 103 Paraformaldehyde (PFA) .................................... 270, 273, 290, 295, 296 Particle ..................................................44, 115, 117, 126, 161, 162, 170, 212, 233–240, 244, 259, 296, 312, 317, 318, 324 Pathogenicity lyme disease ............................................................. 159 swine dysentery ....................................................... 159 syphilis...................................................................... 159 zoonosis leptospirosis ............................................. 159 Peptide chain ................................................................. 299

BACTERIAL Peptidoglycan (PG) layer..................................18, 84, 85, 96, 97, 105, 126, 299, 300, 304, 306, 311, 337, 359 Peptidoglycan-binding (PGB) domain ....................72, 84 Peptidoglycan-binding motif ......................................... 72 Periplasmic region........................................................... 96 Peristaltic pump........................................... 185, 187, 188 Petri dish dual compartment................................. 150, 151, 153 Phenylmethylsulfonyl fluoride (PMSF) .............. 313, 362 ph imaging..................................................................... 134 Phosphate release assay ................................................. 363 Phototaxis negative..........................................256, 259, 261, 262 positive............................................................ 259–261 Pilus (Pili) type IV pili (T4P)..........................183, 249, 255, 299 Pixel ................................................. 39, 49, 91, 203, 204, 228, 233, 236, 239, 240, 242, 284, 316, 317 Plasmid pACTrc ................................................................86, 88 pBAD vector pBAD24............................................79, 80, 86, 89 pBAD33............................................................... 19 pBAD-PomΔplug ........................................ 86, 89, 92 pBBR111 ............................................... 134, 137, 140 pCold4-pomAB-His6 ...................................... 99, 102 pColdI (pCold-I) ................................................66, 96 pET vector pET15b..................................................... 365, 370 pET21b..................................................... 111, 120 pEVOL-pBpF ........................................ 73, 74, 77, 79 pFJ29 .............................................................. 288, 293 pISM2062.2 ............................................................ 355 pITH104 .............................................................19, 21 pKM170-ST ..................................348, 350, 351, 354 pKM310-ST ...........................................348–350, 355 pKY077............................................................. 5, 9, 13 pLysS ....................................................................... 105 pNS1 ............................................................... 288, 293 pRR51............................................................. 288, 292 pTK170-D..............................................348–350, 355 pTSK170 ...................................................... 73, 75, 80 pYC20..................................................................86, 88 pYC109................................................................86, 88 pYC112................................................................86, 88 pYS3....................................................... 73, 74, 77, 79 pYVM132 ............................................................86, 88 Platinum ..............................................300, 302, 304, 305 Plug ............................................................. 36, 84–86, 89, 92, 96, 97, 143, 257, 258 Plunge-freezing device (Plunge freezer)........................ 46 Polyethylene glycol 8000 (PEG8000)................ 186, 193 Poly-L-lysine.............................................. 37, 39, 41, 139

AND

ARCHAEAL MOTILITY Index 399

Polymerase chain reaction (PCR) colony PCR ............................................................. 297 inverse PCR method ...................................... 353, 355 Polymerization ............................................ 360, 363, 366 Polymorphic forms L (left-handed) state ................................................. 44 R (right-handed) state .............................................. 44 Polymorphic supercoiling ............................................... 44 Polypeptide channel ...................................................... 4, 5 Positively charged nanogold................................ 290, 295 Postprocess ................................................................50, 51 Potential energy ............................................................ 172 Powermeter ................................................................... 257 Promotor ......................................................................... 92 Protease inhibitor phenylmethylsulfonyl fluoride (PMSF)......... 362, 369 Protein dynamics............................................................. 57 Protein export ......................................... 4, 17–19, 23, 25 Protein-protein interaction.......................................57, 72 Protein purification ................................... 67, 98, 99, 120 Protein translocation......................................................... 4 Protofilament................................................................... 44 Proton (H+)......................................................83, 95, 109 Proton (H+) channel................................... 83, 85, 90, 92 Proton (H+) flow ........................................................4, 83 Proton motive force (PMF) ...........................3, 4, 17, 19, 27, 83, 278, 284, 378, 379 Proton/protein antiporter ............................................... 4 Purine nucleoside phosphorylase (PNP) ....................363, 364, 366, 367

Q Quick-freeze deep-etch replica..................................... 300

R Radiation damage .................................................. 49, 228 Reactive oxygen species (ROS) .................................... 379 Real-space refinement ..................................................... 51 Refinement three-dimensional (3D) auto-refine......................... 51 Resolution ............................................35, 50, 51, 57, 99, 105, 177, 203, 204, 212, 229, 233, 236, 239, 242–244, 267, 318, 328, 331, 332 Resolution estimation ................................................... 236 Rezasurin ....................................................................... 185

S Sample distribution .............................................. 216, 217 Secretion system rhoptry secretion system......................................... 213 type II secretion system (T2SS) ............................. 212 type III secretion system (T3SS)

BACTERIAL AND ARCHAEAL MOTILITY

400 Index

flagellar protein export apparatus (fT3SS), cytoplasmic ATPase complex...................... 3, 5 flagellar protein export apparatus (fT3SS), transmembrane export gate complex..............3 type IV filament (TFF) ........................................... 183 type IV secretion system cag type IV secretion system ............................ 212 conjugation system ........................................... 212 dot/Icm type IVB secretion system................. 212 type V secretion system........................................... 212 type VI secretion system ......................................... 212 type IX secretion system (T9SS) PorLM ............................................................... 269 T9SS protein ............................................ 268, 278 Semiconductor laser...................................................... 174 Sialylated oligosaccharide ..........311, 312, 321, 327, 328 Signal transduction ....................................................... 255 Simulation molecular dynamics (MD) simulations .............27–30, 32, 58 parallel cascade selection molecular dynamics (PaCS-MD) simulations................................ 31 Simultaneous iterative reconstruction technique (SIRT) .......................................................... 233 Slicer...................................................................... 233, 235 Sodium binding site ........................................................ 95 Sodium 2,2-dimethyl-2-silapentane-5-sulfonate (DSS)........................................................61, 63 Sodium dodecyl sulfate-polyacrylamide gel electrophoresis (SDS-PAGE) ............ 8, 11, 12, 21, 24, 25, 48, 76, 78, 96, 99, 102–104, 116, 122, 189, 192, 314–316, 362, 364, 365 Sodium ion (Na+) .............................................. 4, 95, 109 Sodium motive force.............................................. 95, 109 Software alignframes............................................................... 229 AMBER cpptraj module .................................................... 32 pmemd.cuda module ........................................... 32 tleap module ........................................................ 32 Andor iQ3 ............................................................... 289 Andor solis software....................................... 201, 330 ChimeraX........................................................ 214, 239 ClusCo ....................................................................... 32 Coot .....................................................................46, 51 DigitalMicrograph fast Fourier transform (FFT)...........115, 129, 219 Dynamo adaptive bandpass filtering...................... 214, 235, 236, 239, 243, 244 EMAN2 ................................................................... 243 emClarity ................................................................. 243

EMMA....................................................................... 32 EM menu................................................................. 314 EPU software ............................................................ 46 Etomo Build tomogram ................................................ 232 Coarse alignment .............................................. 232 Fine alignment .................................................. 233 PEET ........................................................ 236, 243 Post-processing ................................................. 234 Pre-processing ................................................... 232 Tomogram Generation ..................................... 233 Tomogram positioning ..................................... 233 Fiji ................................................................. 87, 91, 92 framewatcher ......................................... 229, 231, 243 Gctf v1.06.................................................................. 46 IDL ............................................................................ 37 Igor Pro. software .........................201, 205, 330, 332 ImageJ................................................. 37, 91, 92, 129, 163, 178, 258, 259, 261, 283, 322, 324, 377 IMOD...........................................214, 229, 231, 232, 234–235, 239, 243, 244 Kodec ....................................................................... 113 LAS X software........................................................ 217 M.............................................................................. 243 MATLAB ................................ 37, 235, 237, 239, 243 Microsoft Excel ........................ 87, 88, 129, 163, 178 MODELLER ......................................................46, 51 MotionCor2 ...............................................46, 49, 229 NIS Elements AR........................................... 136, 140 PHENIX ..............................................................46, 51 Prism ............................................................. 87, 89, 91 Protparam ................................................................ 370 PyEMMA................................................................... 32 Python ..................................................................... 243 PyTom ..................................................................... 243 RELION-3.0-β2 ....................................................... 46 SerielEM .................................................................. 214 Sherpa ............................................................. 214, 220 TEM User Interface......................214, 217, 220, 239 UCSF Chimera...........................................46, 51, 241 WARP ...................................................................... 243 Solid surfaces ...................................................71, 95, 160, 169, 249, 268, 287, 311, 321, 327, 337 Solubilization ........................................... 5, 6, 11, 96–98, 112, 114, 121, 318 Sonicator..................................................... 105, 122, 128, 313, 362, 364 Spectrometer .................................. 20, 61, 141, 258, 261 Spectrophotometer ........................................... 6, 7, 9–11, 46, 61, 75–78, 99, 102, 105, 111, 112, 116, 122, 149–151, 185, 186, 257, 259, 269, 279, 280, 313, 314, 361–363, 365, 366, 376, 387

BACTERIAL Spectrum aberration ..................................................... 221 Spectrum focus.............................................................. 221 Spheroplast ......................................................... 18, 22, 25 Spot size........................................................220–223, 239 Spreading protein sprA .......................................................................... 287 sprB .......................................................................... 287 sprC .......................................................................... 287 sprD.......................................................................... 287 sprE .......................................................................... 287 sprF........................................................................... 287 sprT .......................................................................... 287 Square ....................................................... 49, 50, 80, 135, 139, 161, 204, 217–219, 225, 233, 242, 300 Stable isotope .................................................................. 57 Stepwise movement 2-D gaussian function............................................. 334 Kerssemakers analysis ..................................... 333, 334 pairwise analysis.............................................. 333, 334 Stereo array isotope labeling (SAIL)...................... 57, 59, 61, 66, 68 Stoichiometry ............................................................4, 110 Streptavidin.................................................. 199, 202, 207 Superresolution .................................................... 228, 231 Surface hydration ................................................................. 148 translocation ............................................................ 249 Surface layer (S-layer) .........................192, 198, 207, 295 Surfactants ............................................................ 148, 202 Swarmers robust....................................................................... 148 temperate ........................................................ 148, 155 Swarming swarm assay............................................ 149, 155, 156 Swimming swim assay ................................................................ 155 swimming force .............................169, 171, 176, 177 swimming speed ................... 172, 176, 177, 203, 204 Switching ................................................................. 44, 58, 105, 147, 197, 198

T Tag FLAG tag................................................................... 13 HA tag ....................................................................... 13 His tag .................................................................13, 96 Tethered cell cell trajectory ........................................................... 278 helical trajectory .......................................... 35, 36, 99, 125, 183, 197, 249, 277 Thermophile ......................................................... 192, 383

AND

ARCHAEAL MOTILITY Index 401

Thiol group ...............................................................73, 78 Thon ring ...................................................................... 219 Three-dimensional (3D) structure......................... 44, 59, 63, 66, 212 Tilt angle............................................................... 228–231 Tilting schemes bidirectional tilting scheme .................................... 229 Tilt series............................ 212, 222, 224, 228–232, 243 Tilt series acquisition .................213, 217, 218, 230, 242 Tissue culture flask .............................................. 313, 314, 324, 331, 339, 340, 350, 353, 377 Tomogram .................................. 212, 231–238, 243, 244 Tomogram reconstruction..................213, 231, 239, 243 Torque .......................................................... 3, 58, 72, 83, 84, 96, 99, 109, 127, 183, 199, 204, 207, 332, 373 Torque generation ...................72, 96, 97, 109, 125, 279 Total dose ............................................................... 49, 228 Trajectory ................................28, 30–32, 126, 129, 161, 233, 256, 257, 278, 279, 282, 283, 332 Transfer shuttle ............................................................. 216 Transfer station..................................................... 213, 217 Transformation electroporation ........................................................ 348 Transmembrane (TM) ............................. 3–5, 17, 27, 71, 72, 83–85, 92, 95–98, 109, 117, 120, 279 Transposon Tn4001 vector ...................... 347, 349–351, 353–356 Trichloroacetic acid (TCA)................................ 21, 24, 25 Tumble tumble bias .............................................................. 148 Tuning-fork-type viscometer........................................ 177 Turnover ................................................................. 84, 341

U Ultracentrifugation density gradient centrifugation ................................ 19 sucrose density gradient............................... 19, 22, 47 ultracentrifuge .........................................7, 20, 22, 46, 51, 62, 99, 114, 115, 186, 187, 190, 191 Ultraviolet (UV)........................................ 72, 73, 80, 369 Uracil ........................................................... 185, 187, 200 UV lamp ............................................................. 75, 77, 80

V Vacuum ....................... 39, 219, 220, 223, 230, 242, 302 Vaseline ................................................................. 322–324 Viscosity ...................... 71, 172, 177, 189, 207, 332, 376 Visualization ............................................... 121, 152, 203, 216, 257, 261, 281, 282, 300 Vitrification.................................................. 214, 218, 239

BACTERIAL AND ARCHAEAL MOTILITY

402 Index W

X

Waring blender ............................................ 186, 189, 192 Weighted back projection ............................................. 233 Wolfe’s mineral solution ............................................... 135 Wolfe’s vitamin solution ...................................... 135, 136

X-ray X-ray crystallography ..........................................29, 59

Z Zero-loss peak (ZLP)........................................... 220, 230