Channelrhodopsin: Methods and Protocols [1st ed.] 9781071608296, 9781071608302

Table of contents :
Front Matter ....Pages i-xii
Front Matter ....Pages 1-1
Molecular Dynamics Simulations of Channelrhodopsin Chimera, C1C2 (Monika R. VanGordon)....Pages 3-15
Computing Potential of the Mean Force Profiles for Ion Permeation Through Channelrhodopsin Chimera, C1C2 (Chad Priest, Monika R. VanGordon, Caroline Rempe, Mangesh I. Chaudhari, Mark J. Stevens, Steve Rick et al.)....Pages 17-28
Nanodisc Reconstitution of Channelrhodopsins Heterologously Expressed in Pichia pastoris for Biophysical Investigations (Maria Walter, Ramona Schlesinger)....Pages 29-48
Characterizing Channelrhodopsin Channel Properties Via Two-Electrode Voltage Clamp and Kinetic Modeling (Lindsey Prignano, Lauren Herchenroder, Robert E. Dempski)....Pages 49-63
Front Matter ....Pages 65-65
Charge Transport by Light-Activated Rhodopsins Determined by Electrophysiological Recordings (Tamara Hussein, Christian Bamann)....Pages 67-84
Probing Channelrhodopsin Electrical Activity in Algal Cell Populations (Oleg A. Sineshchekov, Elena G. Govorunova, John L. Spudich)....Pages 85-96
Two-Photon Optogenetic Stimulation of Drosophila Neurons (Mehmet Fişek, James M. Jeanne)....Pages 97-108
Probing Synaptic Signaling with Optogenetic Stimulation and Genetically Encoded Calcium Reporters (Gabriel B. Borja, Himali Shroff, Hansini Upadhyay, Pin W. Liu, Valeriya Baru, Yung-Chih Cheng et al.)....Pages 109-134
Optogenetics to Interrogate Neuron-Glia Interactions in Pups and Adults (Chloé Habermacher, Blandine Manot-Saillet, Domiziana Ortolani, Fernando C. Ortiz, María Cecilia Angulo)....Pages 135-149
Chronic Optogenetic Pacing of Human-Induced Pluripotent Stem Cell-Derived Engineered Cardiac Tissues (Marc Dwenger, William J. Kowalski, Hidetoshi Masumoto, Takeichiro Nakane, Bradley B. Keller)....Pages 151-169
Front Matter ....Pages 171-171
Patterned Optogenetic Stimulation Using a DMD Projector (Aanchal Bhatia, Sahil Moza, Upinder S. Bhalla)....Pages 173-188
Selecting Channelrhodopsin Constructs for Optimal Visual Restoration in Differing Light Conditions (Tushar H. Ganjawala, Zhuo-Hua Pan)....Pages 189-199
Recording Channelrhodopsin-Evoked Field Potentials and Startle Responses from Larval Zebrafish (Yagmur Idil Ozdemir, Christina A. Hansen, Mohamed A. Ramy, Eileen L. Troconis, Lauren D. McNeil, Josef G. Trapani)....Pages 201-220
Automated Functional Screening for Modulators of Optogenetically Activated Neural Responses in Living Organisms (Ross C. Lagoy, Eric Larsen, Dirk R. Albrecht)....Pages 221-233
Optogenetic Interrogation of ChR2-Expressing GABAergic Interneurons After Transplantation into the Mouse Brain (Muhammad N. Arshad, Gloster B. Aaron, Janice R. Naegele)....Pages 235-259
Application of Targeting-Optimized Chronos for Stimulation of the Auditory Pathway (Antoine Tarquin Huet, Vladan Rankovic)....Pages 261-285
Channelrhodopsins for Cell-Type Specific Illumination of Cardiac Electrophysiology (Marbely C. Fernández, Ramona A. Kopton, Ana Simon-Chica, Josef Madl, Ingo Hilgendorf, Callum M. Zgierski-Johnston et al.)....Pages 287-307
Optogenetic Control of Cardiac Autonomic Neurons in Transgenic Mice (Angel Moreno, Grant Kowalik, David Mendelowitz, Matthew W. Kay)....Pages 309-321
Dissecting Mechanisms of Motivation within the Nucleus Accumbens Using Optogenetics (Shannon L. Cole, Jeffrey J. Olney)....Pages 323-349
Optogenetic Stimulation of the Central Amygdala Using Channelrhodopsin (Anna S. Knes, Charlotte M. Freeland, Mike J. F. Robinson)....Pages 351-376
Optical Manipulation of Perfused Mouse Heart Expressing Channelrhodopsin-2 in Rhythm Control (Xi Wang, Yue Cheng)....Pages 377-390
Chronic Optogenetic Stimulation in Freely Moving Rodents (Thiago C. Moulin)....Pages 391-401
Back Matter ....Pages 403-405

Citation preview

Methods in Molecular Biology 2191

Robert E. Dempski Editor

Channelrhodopsin Methods and Protocols

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.

Channelrhodopsin Methods and Protocols

Edited by

Robert E. Dempski Department of Chemistry and Biochemistry, Worcester Polytechnic Institute, Worcester, MA, USA

Editor Robert E. Dempski Department of Chemistry and Biochemistry Worcester Polytechnic Institute Worcester, MA, USA

ISSN 1064-3745 ISSN 1940-6029 (electronic) Methods in Molecular Biology ISBN 978-1-0716-0829-6 ISBN 978-1-0716-0830-2 (eBook) https://doi.org/10.1007/978-1-0716-0830-2 © Springer Science+Business Media, LLC, part of Springer Nature 2021 This work is subject to copyright. All rights are reserved 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 Channelrhodopsin-1 and -2 are light-gated ion channels located within the eyespot of the single-cell green algae Chlamydomonas reinhardtii. Activation of channelrhodopsins is the first committed step in phototaxis, the process by which alga move towards light. C. reinhardtii can then use light energy as a starting point to create amino acids and sugars, essential biomolecules. Previously, it had been proposed that light could be used to heterologously control cellular process. The molecular identification of channelrhodopsins in 2002 and 2003 heralded an important step in an emerging field, which would later be named optogenetics. Amazingly, it was quickly shown that these algae light-activated proteins could be functionally expressed in eukaryotic cells such as neurons to control neuronal spiking. Since the first molecular identification of channelrhodopsins, a broad range of tools have been created and new approaches developed to both better understand the molecular determinants of channelrhodopsin function as well as to use these and homologous proteins from a variety of species as tools to better understand physiological processes. More recently, channelrhodopsins have become instrumental as a potential treatment for disease states. This book merges approaches in understanding channelrhodopsin function together with methods addressing how channelrhodopsins can be used to address biomedical questions on a cellular or organismal level. By bringing these diverse methods under one cover, the intention of this edition is to provide a resource for those interested in honing their current expertise as well as potentially branching out into new directions. Worcester, MA, USA

Robert E. Dempski

v

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

PART I

BASIC SCIENCE

1 Molecular Dynamics Simulations of Channelrhodopsin Chimera, C1C2. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Monika R. VanGordon 2 Computing Potential of the Mean Force Profiles for Ion Permeation Through Channelrhodopsin Chimera, C1C2 . . . . . . . . . . . . . . . . . . . . Chad Priest, Monika R. VanGordon, Caroline Rempe, Mangesh I. Chaudhari, Mark J. Stevens, Steve Rick, and Susan B. Rempe 3 Nanodisc Reconstitution of Channelrhodopsins Heterologously Expressed in Pichia pastoris for Biophysical Investigations. . . . . . . . . . . . . . . . . . . . Maria Walter and Ramona Schlesinger 4 Characterizing Channelrhodopsin Channel Properties Via Two-Electrode Voltage Clamp and Kinetic Modeling . . . . . . . . . . . . . . . . . . . . . . . Lindsey Prignano, Lauren Herchenroder, and Robert E. Dempski

PART II

v ix

3

17

29

49

CELLS

5 Charge Transport by Light-Activated Rhodopsins Determined by Electrophysiological Recordings . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 67 Tamara Hussein and Christian Bamann 6 Probing Channelrhodopsin Electrical Activity in Algal Cell Populations . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 85 Oleg A. Sineshchekov, Elena G. Govorunova, and John L. Spudich 7 Two-Photon Optogenetic Stimulation of Drosophila Neurons . . . . . . . . . . . . . . . . 97 Mehmet Fis¸ek and James M. Jeanne 8 Probing Synaptic Signaling with Optogenetic Stimulation and Genetically Encoded Calcium Reporters . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 109 Gabriel B. Borja, Himali Shroff, Hansini Upadhyay, Pin W. Liu, Valeriya Baru, Yung-Chih Cheng, Owen B. McManus, Luis A. Williams, Graham T. Dempsey, and Christopher A. Werley 9 Optogenetics to Interrogate Neuron-Glia Interactions in Pups and Adults . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 135 Chloe´ Habermacher, Blandine Manot-Saillet, Domiziana Ortolani, Fernando C. Ortiz, and Marı´a Cecilia Angulo 10 Chronic Optogenetic Pacing of Human-Induced Pluripotent Stem Cell-Derived Engineered Cardiac Tissues . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 151 Marc Dwenger, William J. Kowalski, Hidetoshi Masumoto, Takeichiro Nakane, and Bradley B. Keller

vii

viii

Contents

PART III 11 12

13

14

15

16

17

18

19

20

21

22

ANIMALS

Patterned Optogenetic Stimulation Using a DMD Projector . . . . . . . . . . . . . . . . . Aanchal Bhatia, Sahil Moza, and Upinder S. Bhalla Selecting Channelrhodopsin Constructs for Optimal Visual Restoration in Differing Light Conditions . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Tushar H. Ganjawala and Zhuo-Hua Pan Recording Channelrhodopsin-Evoked Field Potentials and Startle Responses from Larval Zebrafish . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Yagmur Idil Ozdemir, Christina A. Hansen, Mohamed A. Ramy, Eileen L. Troconis, Lauren D. McNeil, and Josef G. Trapani Automated Functional Screening for Modulators of Optogenetically Activated Neural Responses in Living Organisms . . . . . . . . . . . . . . . . . . . . . . . . . . . Ross C. Lagoy, Eric Larsen, and Dirk R. Albrecht Optogenetic Interrogation of ChR2-Expressing GABAergic Interneurons After Transplantation into the Mouse Brain. . . . . . . . . . . . . . . . . . . . Muhammad N. Arshad, Gloster B. Aaron, and Janice R. Naegele Application of Targeting-Optimized Chronos for Stimulation of the Auditory Pathway . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Antoine Tarquin Huet and Vladan Rankovic Channelrhodopsins for Cell-Type Specific Illumination of Cardiac Electrophysiology . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Marbely C. Ferna´ndez, Ramona A. Kopton, Ana Simon-Chica, Josef Madl, Ingo Hilgendorf, Callum M. Zgierski-Johnston, and Franziska Schneider-Warme Optogenetic Control of Cardiac Autonomic Neurons in Transgenic Mice . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Angel Moreno, Grant Kowalik, David Mendelowitz, and Matthew W. Kay Dissecting Mechanisms of Motivation within the Nucleus Accumbens Using Optogenetics . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Shannon L. Cole and Jeffrey J. Olney Optogenetic Stimulation of the Central Amygdala Using Channelrhodopsin . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Anna S. Knes, Charlotte M. Freeland, and Mike J. F. Robinson Optical Manipulation of Perfused Mouse Heart Expressing Channelrhodopsin-2 in Rhythm Control . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Xi Wang and Yue Cheng Chronic Optogenetic Stimulation in Freely Moving Rodents. . . . . . . . . . . . . . . . . Thiago C. Moulin

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

173

189

201

221

235

261

287

309

323

351

377 391 403

Contributors GLOSTER B. AARON • Department of Biology, Program in Neuroscience and Behavior, Wesleyan University, Middletown, CT, USA DIRK R. ALBRECHT • Department of Biomedical Engineering, Worcester Polytechnic Institute, Worcester, MA, USA; Department of Biology and Biotechnology, Worcester Polytechnic Institute, Worcester, MA, USA MARI´A CECILIA ANGULO • Universite´ de Paris, Institute of Psychiatry and Neuroscience of Paris (IPNP), INSERM U1266, Paris, France; GHU PARIS Psychiatrie & Neurosciences, Paris, France MUHAMMAD N. ARSHAD • Department of Biology, Program in Neuroscience and Behavior, Wesleyan University, Middletown, CT, USA CHRISTIAN BAMANN • Department of Biophysical Chemistry, Max Planck Institute of Biophysics, Frankfurt am Main, Germany VALERIYA BARU • Q-State Biosciences Inc., Cambridge, MA, USA UPINDER S. BHALLA • National Centre for Biological Sciences (NCBS), Tata Institute of Fundamental Research (TIFR), Bangalore, Karnataka, India AANCHAL BHATIA • National Centre for Biological Sciences (NCBS), Tata Institute of Fundamental Research (TIFR), Bangalore, Karnataka, India GABRIEL B. BORJA • Q-State Biosciences Inc., Cambridge, MA, USA MANGESH I. CHAUDHARI • Sandia National Laboratories, Albuquerque, NM, USA YUE CHENG • Department of Cardiology, Renmin Hospital of Wuhan University, Wuhan, People’s Republic of China; Cardiovascular Research Institute, Wuhan University, Wuhan, People’s Republic of China; Hubei Key Laboratory of Cardiology, Wuhan, People’s Republic of China YUNG-CHIH CHENG • Q-State Biosciences Inc., Cambridge, MA, USA SHANNON L. COLE • Department of Anatomy and Neurobiology, University of Maryland, Baltimore, MD, USA GRAHAM T. DEMPSEY • Q-State Biosciences Inc., Cambridge, MA, USA ROBERT E. DEMPSKI • Department of Chemistry and Biochemistry, Worcester Polytechnic Institute, Worcester, MA, USA MARC DWENGER • Kosair Charities Pediatric Heart Research Program, Cardiovascular Innovation Institute, University of Louisville, Louisville, KY, USA; Department of Pharmacology & Toxicology, University of Louisville School of Medicine, Louisville, KY, USA MARBELY C. FERNA´NDEZ • Institute for Experimental Cardiovascular Medicine, University Heart Center Freiburg—Bad Krozingen, Medical Center—University of Freiburg, Freiburg, Germany; Faculty of Medicine, University of Freiburg, Freiburg, Germany; Faculty of Biology, University of Freiburg, Freiburg, Germany MEHMET FIS¸EK • Wolfson Institute for Biomedical Research and Department of Neuroscience, Physiology, and Pharmacology, University College London, London, UK CHARLOTTE M. FREELAND • Neuroscience & Behavior Program, Wesleyan University, Middletown, CT, USA; Department of Biology, Wesleyan University, Middletown, CT, USA

ix

x

Contributors

TUSHAR H. GANJAWALA • Department of Ophthalmology, Visual and Anatomical Sciences (OVAS), Wayne State University School of Medicine, Detroit, MI, USA ELENA G. GOVORUNOVA • McGovern Medical School, The University of Texas Health Science Center at Houston, Houston, TX, USA CHLOE´ HABERMACHER • Universite´ de Paris, Institute of Psychiatry and Neuroscience of Paris (IPNP), INSERM U1266, Paris, France CHRISTINA A. HANSEN • Department of Biology and Neuroscience Program, Amherst College, Amherst, MA, USA LAUREN HERCHENRODER • Department of Chemistry and Biochemistry, Worcester Polytechnic Institute, Worcester, MA, USA INGO HILGENDORF • Faculty of Medicine, University of Freiburg, Freiburg, Germany; Department of Cardiology I, University Heart Center Freiburg—Bad Krozingen, Freiburg, Germany ANTOINE TARQUIN HUET • Institute for Auditory Neuroscience and InnerEarLab, University Medical Center Go¨ttingen, Go¨ttingen, Germany; Auditory Neuroscience and Optogenetics Laboratory, German Primate Center, Go¨ttingen, Germany TAMARA HUSSEIN • Department of Biophysical Chemistry, Max Planck Institute of Biophysics, Frankfurt am Main, Germany JAMES M. JEANNE • Department of Neuroscience, Yale School of Medicine, New Haven, CT, USA MATTHEW W. KAY • Department of Biomedical Engineering, The George Washington University, Washington, DC, USA BRADLEY B. KELLER • Kosair Charities Pediatric Heart Research Program, Cardiovascular Innovation Institute, University of Louisville, Louisville, KY, USA; Department of Pharmacology & Toxicology, University of Louisville School of Medicine, Louisville, KY, USA; Cincinnati Children’s Heart Institute, Cincinnati Children’s Hospital Medical Center, Louisville, KY, USA ANNA S. KNES • Neuroscience & Behavior Program, Wesleyan University, Middletown, CT, USA; Department of Psychology, Wesleyan University, Middletown, CT, USA RAMONA A. KOPTON • Institute for Experimental Cardiovascular Medicine, University Heart Center Freiburg—Bad Krozingen, Medical Center—University of Freiburg, Freiburg, Germany; Faculty of Medicine, University of Freiburg, Freiburg, Germany; Faculty of Biology, University of Freiburg, Freiburg, Germany GRANT KOWALIK • Department of Biomedical Engineering, The George Washington University, Washington, DC, USA WILLIAM J. KOWALSKI • Kosair Charities Pediatric Heart Research Program, Cardiovascular Innovation Institute, University of Louisville, Louisville, KY, USA; Laboratory of Stem Cell and Neuro-Vascular Biology, Cell and Developmental Biology Center, National Heart, Lung, and Blood Institute, NIH, Bethesda, MD, USA ROSS C. LAGOY • Department of Biomedical Engineering, Worcester Polytechnic Institute, Worcester, MA, USA ERIC LARSEN • Department of Biomedical Engineering, Worcester Polytechnic Institute, Worcester, MA, USA PIN W. LIU • Q-State Biosciences Inc., Cambridge, MA, USA JOSEF MADL • Institute for Experimental Cardiovascular Medicine, University Heart Center Freiburg—Bad Krozingen, Medical Center—University of Freiburg, Freiburg, Germany; Faculty of Medicine, University of Freiburg, Freiburg, Germany

Contributors

xi

BLANDINE MANOT-SAILLET • Universite´ de Paris, Institute of Psychiatry and Neuroscience of Paris (IPNP), INSERM U1266, Paris, France HIDETOSHI MASUMOTO • Kosair Charities Pediatric Heart Research Program, Cardiovascular Innovation Institute, University of Louisville, Louisville, KY, USA; Department of Cell Growth and Differentiation, Center for iPS Cell Research and Application, Kyoto University, Kyoto, Japan; Department of CV Surgery, Kyoto University Graduate School of Medicine, Kyoto, Japan; Clinical Translational Research Program, RIKEN Center for Biosystems Dynamics Research, Kobe, Japan OWEN B. MCMANUS • Q-State Biosciences Inc., Cambridge, MA, USA LAUREN D. MCNEIL • Department of Biology and Neuroscience Program, Amherst College, Amherst, MA, USA DAVID MENDELOWITZ • Department of Pharmacology and Physiology, The George Washington University, Washington, DC, USA ANGEL MORENO • Department of Biomedical Engineering, The George Washington University, Washington, DC, USA THIAGO C. MOULIN • Department of Neuroscience, Uppsala University, Uppsala, Sweden; Institute of Medical Biochemistry, Federal University of Rio de Janeiro, Rio de Janeiro, Brazil SAHIL MOZA • National Centre for Biological Sciences (NCBS), Tata Institute of Fundamental Research (TIFR), Bangalore, Karnataka, India JANICE R. NAEGELE • Department of Biology, Program in Neuroscience and Behavior, Wesleyan University, Middletown, CT, USA TAKEICHIRO NAKANE • Kosair Charities Pediatric Heart Research Program, Cardiovascular Innovation Institute, University of Louisville, Louisville, KY, USA; Department of CV Surgery, Kyoto University Graduate School of Medicine, Kyoto, Japan JEFFREY J. OLNEY • Department of Psychology, University of Michigan, Ann Arbor, MI, USA FERNANDO C. ORTIZ • Instituto de Ciencias Biome´dicas, Facultad de Ciencias de la Salud, Universidad Aut
onoma de Chile, Santiago, Chile DOMIZIANA ORTOLANI • Universite´ de Paris, Institute of Psychiatry and Neuroscience of Paris (IPNP), INSERM U1266, Paris, France YAGMUR IDIL OZDEMIR • Department of Biology and Neuroscience Program, Amherst College, Amherst, MA, USA ZHUO-HUA PAN • Department of Ophthalmology, Visual and Anatomical Sciences (OVAS), Wayne State University School of Medicine, Detroit, MI, USA CHAD PRIEST • Sandia National Laboratories, Albuquerque, NM, USA LINDSEY PRIGNANO • Department of Chemistry and Biochemistry, Worcester Polytechnic Institute, Worcester, MA, USA MOHAMED A. RAMY • Department of Biology and Neuroscience Program, Amherst College, Amherst, MA, USA VLADAN RANKOVIC • Institute for Auditory Neuroscience and InnerEarLab, University Medical Center Go¨ttingen, Go¨ttingen, Germany; Restorative Cochlear Genomics Group, Auditory Neuroscience and Optogenetics Laboratory, German Primate Center, Go¨ttingen, Germany CAROLINE REMPE • Sandia National Laboratories, Albuquerque, NM, USA SUSAN B. REMPE • Sandia National Laboratories, Albuquerque, NM, USA STEVE RICK • Department of Chemistry, University of New Orleans, New Orleans, LA, USA

xii

Contributors

MIKE J. F. ROBINSON • Neuroscience & Behavior Program, Wesleyan University, Middletown, CT, USA; Department of Psychology, Wesleyan University, Middletown, CT, USA RAMONA SCHLESINGER • Experimental Physics: Genetic Biophysics, Freie Universit€ a t Berlin, Berlin, Germany FRANZISKA SCHNEIDER-WARME • Institute for Experimental Cardiovascular Medicine, University Heart Center Freiburg—Bad Krozingen, Medical Center—University of Freiburg, Freiburg, Germany; Faculty of Medicine, University of Freiburg, Freiburg, Germany HIMALI SHROFF • Q-State Biosciences Inc., Cambridge, MA, USA ANA SIMON-CHICA • Institute for Experimental Cardiovascular Medicine, University Heart Center Freiburg—Bad Krozingen, Medical Center—University of Freiburg, Freiburg, Germany; Faculty of Medicine, University of Freiburg, Freiburg, Germany OLEG A. SINESHCHEKOV • McGovern Medical School, The University of Texas Health Science Center at Houston, Houston, TX, USA JOHN L. SPUDICH • McGovern Medical School, The University of Texas Health Science Center at Houston, Houston, TX, USA MARK J. STEVENS • Sandia National Laboratories, Albuquerque, NM, USA JOSEF G. TRAPANI • Department of Biology and Neuroscience Program, Amherst College, Amherst, MA, USA EILEEN L. TROCONIS • Department of Biology and Neuroscience Program, Amherst College, Amherst, MA, USA HANSINI UPADHYAY • Q-State Biosciences Inc., Cambridge, MA, USA MONIKA R. VANGORDON • Department of Chemistry, University of New Orleans, New Orleans, LA, USA MARIA WALTER • Experimental Physics: Genetic Biophysics, Freie Universit€ a t Berlin, Berlin, Germany XI WANG • Department of Cardiology, Renmin Hospital of Wuhan University, Wuhan, People’s Republic of China; Cardiovascular Research Institute, Wuhan University, Wuhan, People’s Republic of China; Hubei Key Laboratory of Cardiology, Wuhan, People’s Republic of China CHRISTOPHER A. WERLEY • Q-State Biosciences Inc., Cambridge, MA, USA LUIS A. WILLIAMS • Q-State Biosciences Inc., Cambridge, MA, USA CALLUM M. ZGIERSKI-JOHNSTON • Institute for Experimental Cardiovascular Medicine, University Heart Center Freiburg—Bad Krozingen, Medical Center—University of Freiburg, Freiburg, Germany; Faculty of Medicine, University of Freiburg, Freiburg, Germany

Part I Basic Science

Chapter 1 Molecular Dynamics Simulations of Channelrhodopsin Chimera, C1C2 Monika R. VanGordon Abstract Molecular dynamics (MD) simulations have been successfully used for modeling dynamic behavior of biologically relevant systems, such as ion channels in representative environments to decode protein structure-function relationships. Protocol presented here describes steps for generating input files and modeling a monomer of transmembrane cation channel, channelrhodopsin chimera (C1C2), in representative environment of 1,2-dioleoyl-sn-glycero-3-phosphatidylcholine (DOPC) planar lipid bilayer, TIP3P water and ions (Na+ and Cl
) using molecular dynamics package NAMD, molecular graphics/analysis tool VMD, and other relevant tools. MD simulations of C1C2 were performed at 303.15 K and in constant particle number, isothermal-isobaric (NpT) ensemble. The results of modeling have helped understand how key interactions in the center of the C1C2 channel contribute to channel gating and subsequent solvent transport across the membrane. Key words Molecular dynamics, Transmembrane protein, Channelrhodopsin, Protein simulations, Computational protein modeling

1

Introduction In all-atom, classical molecular dynamics simulations, each modeled atom moves according to the Newtonian equations of motion:   ∂U ! , ! , . . . , ! r1 r2 rn m α´! ¼
, α ¼ 1, 2, . . . , N r ∂! r α

are mass and position of atom α and U is a where mα and ! rα position-dependent potential energy of the system of N atoms and therefore couples the motion of atoms. The potential energy and its parameters (force field) represent bonded (bonds, angles) and non-bonded (van der Waals, Coulombic) interactions among modeled atoms. The force fields are developed for specific combinations of atoms, for instance, each amino acid is described by a distinct set of parameters to reproduce unique biochemical Robert E. Dempski (ed.), Channelrhodopsin: Methods and Protocols, Methods in Molecular Biology, vol. 2191, https://doi.org/10.1007/978-1-0716-0830-2_1, © Springer Science+Business Media, LLC, part of Springer Nature 2021

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Monika R. VanGordon

properties. Consequently, molecular dynamics simulations are time-dependent equilibration of interactions among neighboring atoms, and from MD trajectories, one can quantify fluctuations of structural properties (backbone displacement, distances among functionally relevant residues, number of hydrogen bonds, etc.), all of which correlate with proteins’ dynamic nature and function. Channelrhodopsins (ChRs) are light-gated transmembrane ion channels [1], being actively used to control excitable cells in mice and humans [2, 3], and little is known about the molecular mechanism of the protein’s action. ChRs are originally found in microalgae from the genus Chlamydomonas. Upon light activation, all-trans retinal bound to lysine 292 (C1C2 numbering, [4]) via Schiff base undergoes isomerization to a 13-cis form. Subsequent onset of changes in the intramolecular interaction network results in channel opening, hydration, cation permeation, and cell membrane depolarization [5, 6]. Consequently, the process triggers migration of algae toward the environment optimal for photosynthesis. Atomistic molecular dynamics simulations of channelrhodopsins in physiologically relevant environments have successfully been used to help characterize models of channel gating mechanisms and the influence of intramolecular networks of interactions on channel kinetics [6, 7]. The goal of this work is to present an exemplary workflow for modeling channelrhodopsin in a representative environment of 1,2-dioleoyl-sn-glycero-3-phosphatidylcholine (DOPC) planar lipid bilayer, TIP3P water, and ions (Na+ and Cl
) using molecular dynamics package NAMD, [8] molecular graphics/analysis tool VMD [9], and other relevant tools.

2

Materials A parallel molecular dynamics package Nanoscale Molecular Dynamics (NAMD, formerly Not Another Molecular Dynamics Program) and molecular graphics software Visual Molecular Dynamics (VMD) can be downloaded from the website of Theoretical and Computational Biophysics Group at the University of Illinois Urbana-Champaign (http://www.ks.uiuc.edu/). NAMD has been optimized to utilize hundreds of cores, both CPU and GPU, and thus is generally used in high-performance supercomputing centers as well as in the cloud-computing environment. Following files are required to run NAMD simulations: l

Protein coordinate file in Protein Data Bank (pdb) format with atomic coordinates and records related to protein origin, secondary structure, authors, etc. Here crystal structure of channelrhodopsin chimera, C1C2, PDB accession code 3UG9 was used.

Molecular Dynamics Simulations of Channelrhodopsin Chimera, C1C2

3

5

l

Protein structure file (psf) with atom types and connectivity definitions as described in an associated force field; generation of this file is described in this document; the psf file is coupled with atomic coordinates file (pdb) and topology file which come with the force field parameter package.

l

Molecular dynamics configuration files explained in this document.

l

CHARMM36 all-atom force field parameters (http://mackerell. umaryland.edu/charmm_ff.shtml, [10])—force constants, equilibrium distances, angles, charges, and van der Waals parameters used for calculations of potential energy of simulated system.

Methods

3.1 Protein Coordinate File Preparation

1. From Orientations of Proteins in Membranes database [11] (OPM, https://opm.phar.umich.edu; see Note 1), download file with channelrhodopsin chimera coordinates: PDB ID 3UG9. 2. Open VMD. To upload protein coordinated (3UG9.pdb), go to Extensions > TkConsole and in the console type cd path_to_saved_pdb/ mol new 3UG9.pdb

3. To select a monomer (chain A) and save as pdb file, in TkConsole, type: set prot [atomselect top "chain A and (protein or resname RET)"] $prot writepdb 3UG9-chainA.pdb

Caution: this will remove co-crystallized water molecules. Water atoms can be saved by typing in console: set wat [atomselect top "chain A and water"] $wat writepdb 3UG9-waterA.pdb

4. Load the monomer structure (3UG9-chainA.pdb) to VMD. You will notice that some residues (24–48, 110–117, 327–332, 343–356) are missing from the loop regions. These regions can be rebuilt using, for instance, I-TASSER [12–14], an online protein structure and function prediction server, or other homology/loop modeling tool of choice (see Note 2). 5. For modeling, the top homology model predicted by I-TASSER can be used, or the resolved loop residues can be transferred from a model to crystal structure of a protein’s monomer prepared in previous step (3UG9-chainA.pdb) (see Fig. 1).

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Fig. 1 Simulation snapshot of a cross section of a channelrhodopsin chimera C1C2 (pink α-helices), with all-trans retinal (black sticks) bound to Lys292 in representative environment of lipid bilayer consisting of 200 DOPC molecules (100 per layer, sticks), TIP3P water (blue surface), and ions Na+ (yellow spheres) and Cl
(cyan spheres) added to neutralize protein’s net charge and provide 0.15 M salt concentration 3.2 Simulation Box Preparation

Input simulation box for NAMD simulations can be generated either in VMD or through a web server such as CHARMM-GUI Membrane Builder [15, 16]. The CHARMM-GUI server provides an easy to follow workflow for protein immersion in lipids and addition of solvent. The coordinate (pdb), structure (psf), force field parameter/topology files, and configuration files required for running NAMD are available for download in the final step of CHARMM-GUI workflow. In the CHARMM-GUI (http://www.charmm-gui.org) Input Generator menu, select Membrane Builder, Bilayer Builder. Scroll to the bottom of the page and select Protein/Membrane system. Upload PDB file with the C1C2 coordinates and rebuilt loops, 3UG9-chainA.pdb, and select PDB format. In PDB Manipulation Options, rename RET to LYR. Caution: CHARMM-GUI may not handle non-standard residues such as retinal properly; see Note 3. PDB Info tab allows for elementary coordinate file manipulation. Select terminal group patching to add protein N- and C-termini. Select: model the missing residues, if these have not been rebuilt prior to submitting the coordinates (pdb) file to the CHARMMGUI server. Similarly, select protonation of ionizable residues if that has not been previously resolved. Protein protonation can be assessed based on historical protein knowledge and/or verified

Molecular Dynamics Simulations of Channelrhodopsin Chimera, C1C2

7

through modeling of side chain ionization constants using, e.g., pKa value prediction software PROPKA [17]. Select automatically detected disulfide bonds. To generate a set of inputs for molecular dynamics simulations, complete the following steps: Step 1: The coordinates of protein (3UG9.pdb) have been retrieved from the database of proteins aligned to membrane normal; thus, select “Use PDB Orientation”. Alternatively, for a C1C2 monomer, downloaded from the RSCB Protein Data Bank, an alignment to the first principal axis along Z could be an acceptable option. One can generate pore water and measure pore size using either protein geometry or cylindrical radius of select A˚ngstroms. Step 2: For own records, note calculated cross sectional area. Then, select bilayer composition and size in the system size determination options. To generate planar DOPC lipid bilayer, select heterogeneous lipid box type, rectangular, and water thickness 15 A˚. Select length of XY lipid patch based on numbers of lipid components with XY dimensions ratio equal 1. Select a homogeneous lipid bilayer of 100 DOPC molecules pear each leaflet (see Notes 4–6). Step 3: Record dimensions of the generated simulation box: A 88.9 Dimension along the A (X) axis B 88.9 Dimension along the B (Y) axis C 120.9 Dimension along the C (Z) axis

For optimal building of lipid bilayer around protein, use the replacement method. Select ions (Na+, Cl
) to achieve concentration 0.15 M and neutralize the protein net charge. Step 4: Lipid generation is complete. Continue to build ion and water box. Step 5: Continue to assemble DOPC lipid bilayer membrane, water, and protein components. Step 6: Generate NAMD input for constant particle number, isobaric-isothermal (NpT) simulations. Select automatic generation of grid information for evaluating electrostatic interactions using the particle mesh Ewald (PME; see Note 7) method. Set temperature to 303.15 K. 3.3 Molecular Dynamics Simulations Configuration Parameters

Modeled system equilibration can generally be divided in a minimum of four minimization-equilibration cycles which encompass: 1. Lipid tail relaxation with lipid head, protein, and solvent atoms constrained. 2. System relaxation with protein heavy atoms harmonically ˚ ). restrained (suggested force constant ¼ 1 kcal/mol/A

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Monika R. VanGordon

3. Equilibration with all atoms released. 4. Production run from which system properties are obtained (see Notes 7–10). The equilibration of the protein simulated in a given environment and thus minimum equilibration time can be approximated based on the root-mean square deviation (RMSD) fluctuation equilibration around a constant value. Depending on system, this may require tens to hundreds of nanoseconds of simulations per studied system. The NAMD simulation configuration files can be obtained from CHARMM-GUI workflow.

4

Notes 1. For immersion and simulations of proteins in lipid bilayers, the Orientations of Proteins in Membranes (OPM, https://opm. phar.umich.edu) is a preferred source of protein coordinates. The OPM database contains proteins from the RCSB Protein Data Bank (https://www.rcsb.org) pre-oriented with respect to the hydrophobic core of the lipid bilayer. 2. Crystal structures of proteins often lack amino acid regions. Therefore, before modeling these proteins, it is recommended to either review a structure file header or align the sequence of the structure (pdb) file with the expected sequence (FASTA) file to determine the missing regions. Utilization of homology modeling has been proven successful for rebuilding the regions missing from the protein coordinate files [18, 19]. A comparison or recommendation of the most suitable homology modeling package or software is beyond the scope of this document (exemplary reviews on homology modeling: [20, 21]). 3. It is highly recommended to load generated simulation box in VMD and visually inspect the system. If any unexpected atom binding is detected, e.g., between retinal and the protein, regenerate protein coordinate and structure files using psfgen in VMD. To do that, in the pdb file containing C1C2 monomer coordinates with rebuilt loops, e.g., 3UG9-chainA.pdb, replace: (a) Residue name of lysine 296 (this residue corresponds to lysine that retinal is bound to via Schiff base) from LYS to LYR (b) Retinal residue number to 296 and name RET to LYR Next, at the end of the Charmm36 topology file append retinal topology information from prior works [22–26] available at https://www.ks.uiuc.edu/Research/namd/wiki/ index.cgi?ParameterTopologyRepository. In VMD console,

Molecular Dynamics Simulations of Channelrhodopsin Chimera, C1C2

9

go to directory that contains input coordinates (3UG9-chainA. pdb) and topology file updated with retinal information (top_all36_prot_with_LYR.rtf) files. Then, source do_psf. tcl—a file containing the following instructions: resetpsf package require psfgen topology top_all36_prot_with_LYR.rtf pdbalias residue HIS HSD pdbalias atom ILE CD1 CD pdbalias atom SER HG HG1 pdbalias residue HIS HSD pdbalias atom LYS 1HZ HZ1 pdbalias atom LYS 2HZ HZ2 pdbalias atom LYS 3HZ HZ3 pdbalias atom ARG 1HH1 HH11 pdbalias atom ARG 2HH1 HH12 pdbalias atom ARG 1HH2 HH21 pdbalias atom ARG 2HH2 HH22 pdbalias atom ASN 1HD2 HD21 pdbalias atom ASN 2HD2 HD22 pdbalias atom GLU OT1 OE1 pdbalias atom GLU OT2 OE2 segment C1C2 {pdb 3UG9-chainA.pdb } coordpdb 3UG9-chainA.pdb C1C2 regenerate angles dihedrals guesscoord writepdb prot.pdb writepsf prot.psf mol new prot.psf mol addfile prot.pdb

The resulting protein coordinate/structure file pair can be merged with the environment files built in, e.g., CHARMMGUI in the following sequence of commands typed in VMD console: mol new prot_lipid_solvent.psf ;# complete structure file generated in CHARMM-GUI mol addfile prot_lipid_solvent.pdb ;# complete coordinate file generated in CHARMM-GUI

10

Monika R. VanGordon set notprot [atomselect top "not protein"] ;# select all but protein $notprot writepsf not_protein.psf ;# save all but protein $notprot writepdb not_protein.pdb

Next the lipid and solvent files (not_protein.psf/pdb) can be combined with rebuilt protein files (prot.psf/pdb) using VMD > VMD Main > Extensions > Modeling > Merge Files application. 4. Based on author’s observation, 1-palmitoyl-2-oleoyl-sn-glycero-3-phosphatidylcholine (POPC) is a lipid commonly used in simulations of proteins in representative environments. However, depending on scientific question at hand, it may be beneficial to use other lipids (such as DOPC as described here and in [27]) or physiologically relevant heterogeneous mixtures. Most commonly, simulations of protein’s dynamical properties are conducted in lipids with melting temperatures below the selected simulation temperature. 5. The size of simulation box for protein modeling with periodic boundary conditions should be large enough to encompass entirety of protein and assure that the protein is separated from its periodic image by at least the distance chosen for calculations of non-bonded interactions. It is recommended, especially in case of proteins with extended loops, termini, or other flexible regions to monitor the distance of protein from the edge of simulation box and re-starting simulations with corrected size of the simulation box (increased amount of solvent and/or lipids). 6. The non-bonded, electrostatic interactions using the particle mesh Ewald algorithm are typically calculated with a real-space cut-off distance of 14 A˚ and grid width of 1 A˚. The suggested switching distance for non-bonded electrostatics and van der ˚. Waals interactions is 12 A 7. System preparation is a key step in assuring accuracy of the simulated model. Therefore, the relaxation-minimization cycles preceding the production simulation run are always recommended. 8. Properties of the system should not be calculated from a single molecular dynamics simulation frame. It is acceptable to report statistical averages of systems properties calculated from equilibrated production runs or apply clustering algorithms to reduce space of representative frames. Most common properties may include: distances among functionally important residues, backbone RMSD, RMSF, solvent accessible surface area, pKa values (PROPKA package [17]), etc. Most of these

Molecular Dynamics Simulations of Channelrhodopsin Chimera, C1C2

11

properties can be calculated using VMD or associated scripts. Useful tools can also be incorporated in R and Python codes, via utilization of packages such as Bio3D (http://thegrantlab. org/bio3d/index.php), MDAnalysis (https://www. mdanalysis.org), or MDTraj (http://mdtraj.org/1.9.3/). Overview of the analysis methods is beyond the scope of this document [28]. 9. Representative frames can be selected for visualization but justification of the selection should be provided. CatDCD (https://www.ks.uiuc.edu/Development/MDTools/catdcd/ ) is a useful tool for manipulating binary trajectory files. For instance, to extract first ten frames from prod.dcd trajectory file install CatDCD and in terminal type ./catdcd -o frame1-10.dcd -first 1 -last 10 prod.dcd

10. Exemplary NAMD simulations configuration file (run.conf) is shown below. To run NAMD on a non-parallel (non-MPI) workstation, type namd2 run.conf ################################################################### ## JOB DESCRIPTION ## Comments begin with # or ;# ################################################################### # Exemplary configuration file for # equilibration of channelrhodopsin chimera, C1C2 # embedded in membrane, ions and water. # No atom constraints set. Use PME, constant NpT. ################################################################### ## ADJUSTABLE PARAMETERS ## ################################################################### structure 3UG9-chainA.psf ;#generated in CHARMM-GUI and/or psfgen coordinates 3UG9-chainA.pdb ;#coordinates of C1C2 monomer outputName prod01 ;# simulation output file name set temperature 300.15 ;# simulation temperature # Continuing a job from the restart files if {0} { ;# 0 if the section enclosed in {} is not to be executed ;# 1 if this section should be executed, that is when ;# restarting simulations from prot01.restart file set inputname

prod01.restart

binCoordinates

$inputname.coor

;# prefix of restart file ;# restart coordinate file

binVelocities

$inputname.vel

;# remove the "temperature" entry

when velocity file already exist that is when restarting simulation sextendedSystem restart }

$inputname.xsc

;# extended file to be used for

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Monika R. VanGordon

firsttimestep

0

;# 0 if this is the first simulation ;# if simulations restarted, set this to number ;# of simulation steps from previous run +1

##################################################################### ## SIMULATION PARAMETERS ## Charmm force field parameters used here. ## Multiple parameter files accepted ##################################################################### # Input paraTypeCharmm

on

parameters

par_all36_prot_with_LYR.prm ;# protein Charmm

;# if on, input parameters are in Charmm format

parameter file with appended retinal parameters parameters

par_all36_lipid.prm ;# lipid Charmm parameter file

# NOTE: Do not set (comment out) temperature below if you # have also specified a .vel restart file! temperature $temperature # Periodic Boundary Conditions # NOTE: Do not set the periodic cell basis if you have also # specified an .xsc restart file! ;# to get size of the box, load simulation box in VMD, in console type: ;# set wat [atomselect top water] ;# measure minmax $wat ;# to visually check periodic box dimensions, in console type ;# molinfo top set a 90 ;# molinfo top set b 90 ;# molinfo top set c 121 ;# visually inspect periodic boundaries: select VMD Main > Graphics > Representations > Periodic and choose periodic images in x- and ydirections; gap between neighboring images should be negligible. if {1} {

;# use the following size of simulation box

cellBasisVector1 90.0 0.0 0.0 cellBasisVector2 0.0 90.0 0.0 cellBasisVector3 0.0 0.0 121.0 cellOrigin

0.0 0.0 0.0 ;# center of the simulation box

} wrapWater

on

wrapAll

on

# Force-Field Parameters exclude

scaled1-4

1-4scaling

1.0

;# scale interactions between atoms 1 and 4

cutoff

12. ;# van der Waals (vdW) interaction truncation distances

witching

on

;# smooth switching function to truncate the van der

Waals energy smoothly at the cutoff value listed above

Molecular Dynamics Simulations of Channelrhodopsin Chimera, C1C2 switchdist

10. ;# distance at which the vdW energy smoothly

transitions toward 0 at the cutoff distance pairlistdist 14. ;# cutoff used in nonbonded interaction calculations # Integrator Parameters timestep

1.0

rigidBonds

water ;# rigid H-O and H-H distances in water molecules

nonbondedFreq

;# 1 fs per simulation step 1

fullElectFrequency 2 stepspercycle

20

# Set Particle Mesh Ewald (PME) grid size for calculations of # electrostatic interactions # grid size should correspond to at least one grid point per Angstrom # in each direction; grid point number is a multiple of 2, 3, or 5 if {1} {

;# use PMA for electrostatics

PME

yes

PMEGridSpacing 1.0 ;# maximum space between grid points, 1.0 A is recommended } # Constant Temperature Control langevin

on

;# do Langevin dynamics

langevinDamping 1 langevinTemp

;# damping coefficient (gamma) of 5/ps

$temperature

# Constant Pressure Control (variable volume) if {1} { useGroupPressure

yes ;# needed for 2fs steps

useFlexibleCell

yes ;# no for water box, yes for membrane

useConstantArea

no ;# no for water box, yes for membrane

langevinPiston

on

langevinPistonTarget

1.01325 ;# in bar -> 1 atm

langevinPistonPeriod

200.

langevinPistonDecay

50.

langevinPistonTemp

$temperature

} # Set frequency of saving restartfreq

1000 ;# 1000steps = every 1 ps

dcdfreq

1000 ;# simulation frame

xstFreq

1000 ;# extended system trajectory file, ;# contains periodic cell parameters

outputEnergies 50

;# energy output frequency

outputPressure 50

;# pressure output frequency

# Define fixed atoms file # fixed.pdb is coordinate file with beta-column values set # to 1 for atoms that should be constrained/fixed during simulations

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Monika R. VanGordon

if {0} { ;# change to 1 if atoms indicated in fixed.pdb should be constrained/fixed during simulations fixedAtoms

on

fixedAtomsFile

fixed.pdb

fixedAtomsCol

B

fixedAtomsForces on } ################################################################ ## Run molecular dynamics ################################################################ # Number of minimization steps if {1} { minimize

2000

reinitvels

$temperature

} run 5000000 ;# simulation time of 5 ns

References 1. Nagel G, Szellas T, Huhn W et al (2003) Channelrhodopsin-2, a directly light-gated cation-selective membrane channel. Proc Natl Acad Sci U S A 100:13940–13945. https:// doi.org/10.1073/pnas.1936192100 2. Boyden E, Zhang F, Bamberg E et al (2005) Millisecond-timescale, genetically targeted optical control of neural activity. Nat Neurosci 8:1263–1268 3. Bruegmann T, Malan D, Hesse M et al (2010) Optogenetic control of heart muscle in vitro and in vivo. Nat Methods 7:897–900 4. Kato HE, Zhang F, Yizhar O et al (2012) Crystal structure of the channelrhodopsin light-gated cation channel. Nature 482:369–374. https://doi.org/10.1038/ nature10870 5. Ardevol A, Hummer G (2018) Retinal isomerization and water-pore formation in channelrhodopsin-2. Proc Natl Acad Sci 115:3557–3562 6. Kuhne J, Eisenhauer K, Ritter E et al (2015) Early formation of the ion-conducting pore in channelrhodopsin-2. Angew Chem Int Ed 54:4953–4957. https://doi.org/10.1002/ anie.201410180 7. Mu¨ller M, Bamann C, Bamberg E, Ku¨hlbrandt W (2015) Light-induced helix movements in channelrhodopsin-2. J Mol Biol 427:341–349. https://doi.org/10.1016/j.jmb.2014.11.004

8. Phillips JC, Braun R, Wang W et al (2005) Scalable molecular dynamics with NAMD. J Comput Chem 26:1781–1802. https://doi. org/10.1002/jcc.20289 9. Humphrey W, Dalke A, Schulten K (1996) VMD—visual molecular dynamics. J Molec Graphics 14:33–38 10. Huang J, MacKerell AD (2013) CHARMM36 all-atom additive protein force field: validation based on comparison to NMR data. J Comput Chem 34:2135–2145. https://doi.org/10. 1002/jcc.23354 11. Lomize MA, Lomize AL, Pogozheva ID, Mosberg HI (2006) OPM: orientations of proteins in membranes database. Bioinformatics 22:623–625 12. Yang J, Yan R, Roy A et al (2015) The I-TASSER suite: protein structure and function prediction. Nat Methods 12:7–8. https://doi. org/10.1038/nmeth.3213 13. Roy A, Kucukural A, Zhang Y (2010) I-TASSER: a unified platform for automated protein structure and function prediction. Nat Protoc 5:725–738. https://doi.org/10.1038/ nprot.2010.5 14. Zhang Y (2008) I-TASSER server for protein 3D structure prediction. BMC Bioinformatics 9:40. https://doi.org/10.1186/1471-21059-40

Molecular Dynamics Simulations of Channelrhodopsin Chimera, C1C2 15. Jo S, Kim T, Iyer VG, Im W (2008) CHARMM-GUI: a web-based graphical user interface for CHARMM. J Comput Chem 29:1859–1865. https://doi.org/10.1002/ jcc.20945 16. Jo S, Lim JB, Klauda JB, Im W (2009) CHARMM-GUI membrane builder for mixed bilayers and its application to yeast membranes. Biophys J 97:50–58. https://doi. org/10.1016/j.bpj.2009.04.013 17. Olsson MHM, Søndergaard CR, Rostkowski M, Jensen JH (2011) PROPKA3: consistent treatment of internal and surface residues in empirical pKa predictions. J Chem Theory Comput 7:525–537. https://doi.org/ 10.1021/ct100578z 18. Jamroz M, Kolinski A (2010) Modeling of loops in proteins: a multi-method approach. BMC Struct Biol 10:5 19. van Beusekom B, Joosten K, Hekkelman ML et al (2018) Homology-based loop modeling yields more complete crystallographic protein structures. Int Union Crystallogr 5:585–594 20. Wallner B, Elofsson A (2005) All are not equal: a benchmark of different homology modeling programs. Protein Sci 15:1315–1327 21. Muhammed MT, Aki-Yalcin E (2019) Homology modeling in drug discovery: overview, current applications, and future perspectives. Chem Biol Drug Des 93:12–20 22. Tajkhorshid E, Paizs B, Suhai S (1997) Conformational effects on the proton affinity of the Schiff base in bacteriorhodopsin: a density

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functional study. J Phys Chem B 101:8021–8028. https://doi.org/10.1021/ jp971283t 23. Tajkhorshid E, Suhai S (1999) Influence of the methyl groups on the structure, charge distribution, and proton affinity of the retinal Schiff base. J Phys Chem B 103:5581–5590. https:// doi.org/10.1021/jp983742b 24. Tajkhorshid E, Baudry J, Schulten K, Suhai S (2000) Molecular dynamics study of the nature and origin of Retinal’s twisted structure in bacteriorhodopsin. Biophys J 78:683–693. https://doi.org/10.1016/S0006-3495(00) 76626-4 25. Nina M, Roux B, Smith JC (1995) Functional interactions in bacteriorhodopsin: a theoretical analysis of retinal hydrogen bonding with water. Biophys J 68:25–39. https://doi.org/ 10.1016/S0006-3495(95)80184-0 26. Baudry J, Crouzy S, Roux B, Smith JC (1997) Quantum chemical and free energy simulation analysis of retinal conformational energetics. J Chem Inf Comput Sci 37:1018–1024. https:// doi.org/10.1021/ci9702398 27. VanGordon M, Gyawali G, Rick SW, Rempe SB (2017) Atomistic study of intramolecular interactions in the closed-state channelrhodopsin chimera, C1C2. Biophys J 112:943–952 28. Likhachev IV, Balabaev NK, Galzitskaya OV (2016) Available instruments for analyzing molecular dynamics trajectories. Open Biochem J 10:1–11

Chapter 2 Computing Potential of the Mean Force Profiles for Ion Permeation Through Channelrhodopsin Chimera, C1C2 Chad Priest, Monika R. VanGordon, Caroline Rempe, Mangesh I. Chaudhari, Mark J. Stevens, Steve Rick, and Susan B. Rempe Abstract Umbrella sampling, coupled with a weighted histogram analysis method (US-WHAM), can be used to construct potentials of mean force (PMFs) for studying the complex ion permeation pathways of membrane transport proteins. Despite the widespread use of US-WHAM, obtaining a physically meaningful PMF can be challenging. Here, we provide a protocol to resolve that issue. Then, we apply that protocol to compute a meaningful PMF for sodium ion permeation through channelrhodopsin chimera, C1C2, for illustration. Key words Umbrella sampling (US), Molecular dynamics, Potential of mean force (PMF), Statistical mechanics, Free energies, Reaction coordinate (RC), Weighted histogram method (WHAM), Nanotechnology, Optogenetics

1

Introduction Studying the structure and function of biological systems is relevant to advancing drug design [1–4], bioinspired materials [5–8], and biotechnology [9–11]. These systems, including enzyme-based cancer drugs [3, 4] and enzyme-embedded separation membranes [7, 8], can be cumbersome and costly to handle in laboratory settings. A computational approach can complement wet-lab experiments by providing molecular insight into critical details about how the internal and external protein environments govern a biological process. With this additional knowledge, wet-lab experiments can focus on a small list of cancer drug candidates or membrane pore chemistries rather than running large, costly screening tests. Exploring fundamental molecular properties with computational models can be challenging due to the innate complexity of proteins and the timescales of biological processes. As an example, protein folding can extend beyond millisecond timescales, which is

Robert E. Dempski (ed.), Channelrhodopsin: Methods and Protocols, Methods in Molecular Biology, vol. 2191, https://doi.org/10.1007/978-1-0716-0830-2_2, © Springer Science+Business Media, LLC, part of Springer Nature 2021

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Chad Priest et al.

demanding for molecular simulation. To circumvent challenges of large timescale limitations and adequate phase space sampling, statistical mechanics provides a mathematical tool that permits the study of protein structure and function independent of time. Statistical mechanics assists computational chemists in predicting energies along a reaction pathway. Our focus here is on free energy along a reaction coordinate, in particular the potential of mean force (PMF) for ion permeation through a channel protein. A case study is the channelrhodopsin (ChR) family of proteins, which are naturally occurring membrane transport proteins found as aggregates in eyespot regions of the plasma membrane of algae [12–18]. ChR proteins are activated by incident light that opens the channel gate and permits conduction of mono- and divalent cations down an induced electrochemical gradient. The first ChR structure solved was for a chimera called C1C2 [14]. The channelrhodopsin chimera C1C2 is a 342-residue monomer that consists of an engineered combination of helices I–V from ChR1 and VI–VII from ChR2 [12, 14, 15]. The putative pore is localized among helices I–III and VII. Helix II is lined with acidic glutamic acid residues E90, E97, E101, and E123 that facilitate ion selectivity and conductance (using the ChR2 numbering scheme) [12, 13]. The ion channel protein opens to enable cation conduction across the plasma membrane. Pore opening is initiated by a conformational change in the retinal molecule, from all trans to 13-cis [12, 14–16]. A few computational studies have investigated the hydration properties within the closed and open states of the ChR protein pore and the ion permeation path through the hydrated pore [12, 16–19]. However, the precise mechanism of ion transport through ChR has not been determined. Computational methods can be used to calculate PMFs to provide insight into the molecular details of ion transport, such as which amino acids are involved in catalyzing ion transport. Several computational methods are available for studying the changes in free energy for ions positioned at different locations along a protein permeation pathway. These methods include thermodynamic integration [20, 21], free-energy perturbation [22], metadynamics [23], and quasi-chemical theory [24, 25]. In addition, umbrella sampling (US) is a well-documented, accepted method to determine PMFs and free energy [26]. US is a sampling method that uses a biasing potential to model a chemical reaction in a phase space, from one chemical thermodynamic state to another, along a chosen reaction coordinate. Here, we choose US to compute a PMF because of its wide-spread use for biological systems. Efficient conformational sampling of phase space along a reaction coordinate is needed to compute a PMF, but may be hindered by large energy barriers relative to thermal energies. To circumvent such challenges, application of a harmonic (quadratic) biasing potential can ensure a system experiences the different

Computing Potential of the Mean Force Profiles for Ion Permeation Through. . .

19

thermodynamic states, even for rare events. The biasing potential enables efficient sampling of phase space along the whole reaction coordinate. The quadratic potential produces a parabolic well that resembles the shape of umbrellas. Performing US simulations at targeted positions along the reaction coordinate, by keeping the system localized to a region with a biasing potential, results in a distribution of conformations around that target value, and a histogram is constructed. US collects a biased distribution used to calculate the biased probability of the event Pb(r). After performing US, a post-processing algorithm is used to combine histogram distributions of the reaction coordinate from each window obtained from the series of US simulations. After achieving overlap of neighboring histograms from US, the PMF is “stitched” together with a weighting histogram method (WHAM) [27]. WHAM estimates the unbiased probability distribution (Pub(r)), from a series of biased simulations performed by US, from which the free energy can be computed. WHAM with US has been applied to the study of membrane transport proteins. As an example, molecular details on the energy landscape of activation gating and the mechanism of potassium ion permeation in the bacterial potassium ion channel, KcsA, have been explored with US-WHAM [28–30]. The insights gained from calculation of PMFs may aid the development of therapeutics [2, 31] or bioinspired materials such as membranes to separate specific ions or molecules [6, 7, 11]. Here, we provide a succinct conceptual guide for constructing a PMF for ion transport through proteins, using channelrhodopsin C1C2 chimera as a case study. This guide provides a step-by-step approach for choosing an initial configuration, parameterizing US-WHAM, and investigating histogram overlap. Finally, we discuss the quality of the constructed PMF.

2

Computational Materials We adopt a model of channelrhodopsin chimera, C1C2, based on previously published work that equilibrated the protein in the open state in a lipid bilayer [12, 16]. The closed state structure [14] was utilized (PDB ID:3UG9) to determine the open state structure.

2.1

Simulation Setup

1. The CHARMM-GUI software [32] was used to create a simulation box containing a C1C2 molecule initially in the closed state and immersed in lipid bilayers consisting of ~200 molecules of di-oleoyl-sn-glycero-3-phosphatidylcholine (DOPC) (see Note 1). 2. Molecular dynamics (MD) simulations of the C1C2 chimera in the lipid environment were created with the CHARMM36

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Chad Priest et al.

force field for protein, ions, and lipids, along with the complementary TIP3P water model, while the retinal force field was borrowed from previous works [33–38]. 3. The devised system was optimized and equilibrated. A production simulation was performed with a constant-particle-number and isothermal-isobaric ensemble (NPT), which was implemented with open source NAMD code [39] (see Note 2). 4. The WHAM code devised by Grossfield was used to construct a PMF for the permeation of a sodium ion through the channelrhodopsin chimera, C1C2 [40]. In brief, the reaction coordinate was chosen as a collective variable consisting of a single sodium ion and atoms of the protein (see Note 3).

3

Computational Methods Our strategy for constructing a PMF for cation permeation through the C1C2 ion channel has been outlined. Next, we will apply the outlined steps and discuss the results. Although US-WHAM has been applied effectively, two challenges accompany its use: (1) US-WHAM can yield a PMF curve that lacks physical meaning due to lack of converged results; (2) the detailed approach is not exactly the same system-to-system due to differences in chemical environments. To avoid those problems, we recommend this general strategy for constructing a PMF for cation permeation through a protein ion channel, such as channelrhodopsin chimera, C1C2: 1. Select an initial structural model that represents the reaction path under consideration (see Note 4). 2. Choose the reaction coordinate (RC). We chose a collective variable that represents overall ion permeation along the zdirection of the protein, from one side of the membrane to the other. Positioning a nonphysical dummy atom at the extracellular end of the permeation path, near the ion in its initial configuration, assisted in defining the detailed curvilinear collective variable for the reaction coordinate (see Notes 5–7). 3. Select a force constant. US-WHAM is a histogram method that depends on overlap between the neighboring histograms. This overlap can be controlled by the strength of the force constant (see Note 8). 4. Inspect histogram overlap to ensure a reasonable selection of force constant has been made (see Note 9). 5. Visualize the ion’s progress along the reaction coordinate and relate it to the PMF (see Note 10). 6. Analyze PMF (see Notes 11–13).

Computing Potential of the Mean Force Profiles for Ion Permeation Through. . .

4

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Notes 1. The system contained protein, lipid bilayers, water, and ions (Na+ and Cl
) to neutralize the protein’s net charge and provide 0.15 M salt concentration. The total number of atoms in the system was approximately 96,000. 2. A time step of 1 fs was set for each simulation. The Langevin piston method was used to maintain a constant pressure of 1 atm [41]. The temperature, set to 300 K, was controlled using Langevin dynamics with a coupling coefficient of 1 ps. System equilibration was divided into steps I–IV, each followed by 1000 energy minimization steps. Step I involved relaxation of the lipid tails with protein, lipid head-groups, and ions fixed (0.5 ns). Step II consisted of relaxation of the system with protein heavy atoms harmonically restrained (1 kcal/mol-A˚, duration 1 ns). Step III consisted of equilibration with protein constraints released (100 ns). Step IV comprised the production runs (an additional 20 ns of simulation). 3. We constructed the PMF with WHAM using a tolerance of 10
5 and a bin resolution of 0.2 A˚. 4. Many possibilities can be explored for the initial configuration (Fig. 1). We chose to incorporate the ion within the protein between helices II and VII and closer to the extracellular side for several reasons (Fig. 1b). First, this position allows facile incorporation of the ion into the protein in comparison with pulling the ion into the protein from bulk water. Second, this position ensures the protein is equilibrated with an ion inside the channel. Furthermore, the sodium ion migrates from the extracellular to intracellular side of the membrane in the native system, making the extracellular side a reasonable initial position for the ion (Fig. 1b). 5. An RC may be defined by a collective variable that represents progress along a reaction pathway—in this case, sodium (Na+) ion transports through the C1C2 protein channel. A collective variable is a differentiable function of atomic Cartesian coordinates. Our RC utilizes a one-dimensional collective variable that represents progress along the z-axis in three-dimensional space to model the transport of a single Na+ ion (Fig. 1). 6. Initial choice of center-of-mass as RC resulted in the trapping of Na+ in the middle of the protein, leading to structural change of the protein along with a high energy barrier. The high energy barrier arose due to the direct interaction of the ion with the retinal molecule buried in a narrow pocket at the center of the protein. An alternative choice of RC as Na+ position along the protein permeation pathway resulted in a PMF with no high energy barriers, as expected for a permeating ion.

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Fig. 1 (a) Snapshot from molecular dynamics simulations of open-state channelrhodopsin chimera model, C1C2, in DOPC lipid bilayer and surrounded by 0.15 M NaCl in water. The vertical direction aligns with the zaxis utilized here for PMF calculations. (b) Open-state C1C2 with helices I–VII labeled and retinal shown as atoms. The large blue sphere is a sodium ion, positioned within the permeation pathway between residues I, II, III, and VII [12, 16–18]

7. To progress along the RC and generate initial configurations, the ion was pulled along the z-axis of the protein (Fig. 1b), starting from the center of the protein, and guided to the edge of the channel by a dummy atom. A force constant of 10 kcal/ ˚ 2 was implemented for pulling the Na+ ion through the mol-A ˚ , divided into pathway. The reaction coordinate spans 35 A ˚ 0.5 A intervals. Each interval along the reaction coordinate was sampled for 1 ns of simulation time to ensure equilibration of the system. The last 250 ps interval of each simulation was collected for analysis of the PMF. 8. The force constant, k, plays an important role in determining the quality of the calculated PMF. Constructing a PMF with WHAM from a series of windows depends sensitively on histogram overlap. The strength of the force constant influences histogram height, width, and the overall overlap between two neighboring histograms, which is important for WHAM convergence. Discontinuity, or gaps between neighboring histograms, can produce an unreliable PMF with WHAM. Importantly, the choice of a force constant influences the phase space sampling along a reaction coordinate. Most often, a harmonic bias of strength k is applied, where k is the

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force constant, ro is the position (target window) on the reaction path, and rref,i is the simulated position at that target reference window. The value of k (force constant) must be chosen so that the histogram widths overlap with neighboring histograms along a RC and the ion can cross energy barriers [42, 43]. Larger k may be required for an ion to cross an energy barrier. At the same time, a smaller (weaker force constant) k will yield a wider histogram of conformations, with smaller height for windows along the reaction coordinate. In contrast, a larger (stronger force constant) k will result in a narrower histogram distribution with higher peaks. Narrow distributions combined with a small number of windows will create poor histogram overlap, while wider distributions may miss states along a permeation pathway. 9. While deciding the strength of the force constant, it is important to be cognizant of the window spacing. Performing shorter timescale simulations for many windows leads to better histogram overlap and reduces statistical error compared with longer timescale simulations carried out for fewer windows [42, 43]. In our system, we chose parameters appropriate for computing the PMF for an ion permeation pathway. First, we chose a 10 kcal/mol k constant because, in general, the free energy of an ion permeation pathway does not exceed that value. Thus, a 10 kcal/mol force constant should be strong enough to overcome barriers along the permeation path. A 0.5 A˚ spacing with 1 ns simulation time ensured adequate overlap of the histograms for short simulations along many windows (Fig. 2). w i ðr Þ ¼ k=2 r o
r ref ,i


2

ð1Þ

10. We provide a python script to assist readers in performing a histogram analysis (Fig. 2). This script reads in all histogram datasets, labeled “window_∗,” from individual simulations and globally plots all datasets onto one plot (Fig. 2a). The xaxis is the reaction coordinate and the y-axis gives the counts. The Matplotlib module allows easy on-the-fly analysis of the histograms by zooming in along a series of histograms along the x-axis (Fig. 2b, c). The gaps in neighboring histograms can be a consequence of attractions along the permeation path. Since WHAM depends on overlap of histograms, methods for rectifying poor overlap need to be considered (Fig. 2c). These methods include adding more windows within the region of poor overlap, adjusting the k, or rerunning the simulation from a different starting configuration.

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Fig. 2 Analyzing histogram window overlap by plotting raw histograms with a python script. Plots of (a) all histograms along the reaction coordinate, (b) zoomed distributions between 10 and 16 A˚, and (c) zoomed distributions between 24 and 30 A˚

11. Once the appropriate overlap has been achieved, the overall trajectory along the reaction path can be assessed with a visualizing software, such as VMD [44]. The overall trajectory can be inspected visually to see if the RC reflects a sensible ion transport pathway through the protein. From quick inspection of the current results, the overall ion permeation pathway passes through the channel between helices I, II, and VII, as suggested in the literature [13, 17, 18] (Fig. 3). Another feature observed is that the ion enters from the extracellular side between helices I and VII, permeates between II and VII, and then exits between helices I and VII. 12. We assess whether the PMF makes sense relative to experimental observations. The ion permeates with relatively low free energy barriers, as expected for a channel known to be

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Fig. 3 Permeation pathway of a Na+ (in blue) through channelrhodopsin chimera, C1C2 (yellow), showing (a) a side view and (b) a top view. C1C2 embedded in DOPC membrane, shown in transparent gray and red

permeable to ions (Fig. 4a). Wells and peaks are well-defined, and no extreme energy barriers exceeding the chosen force constant exist. The Na+ ion is overall more stable inside the protein than in bulk water. The interfacial energy between bulk water and protein is close as Na+ enters from bulk water and exits into bulk water. If poor histogram overlap were exhibited, WHAM could yield exaggerated barriers during the stitching process. 13. To further interpret the results, we note that the PMF contains a global minimum at about 30 A˚ (Fig. 4a). This region is the binding pocket where Na+ has the most favorable thermodynamic stability within the protein. In this region, Na+ binds to amino acids glutamate (E90) and asparagine (N258) (Fig. 4b). Interestingly, this site has been suggested to be involved in the closing process of the protein through hydrogen bonding with water [12]. Furthermore, hydration studies suggested that this site is part of the ion conduction pathway [17, 18]. The current simulation work adds to the story. Once retinal experiences its conformational change (from all trans to 13-cis) upon exposure to incident light, the channel opens. This allows Na+ ion to move within the protein, driven in part by Coulombic interactions with charged amino acids. The attractive and repulsive ion interactions with the protein are illustrated by the hills and valleys in the PMF (Fig. 4a).

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Fig. 4 (a) Potential of mean force (PMF) for the permeation of Na+ through the C1C2 channelrhodopsin chimera. (b) Simulation snapshot of the ion location in the C1C2 region corresponding to the minimum in the PMF profile (z ~ 30 A˚). Waters were removed for clarity

Acknowledgments This work was performed, in part, at the Center for Integrated Nanotechnologies, an Office of Science User Facility operated for the U.S. Department of Energy (DOE) Office of Science. Sandia National Laboratories is a multi-mission laboratory managed and operated by National Technology & Engineering Solutions of Sandia, LLC, a wholly owned subsidiary of Honeywell International, Inc., for the U.S. DOE’s National Nuclear Security Administration under contract DE-NA-0003525. The views expressed in the article do not necessarily represent the views of the U.S. DOE or the US government. The authors have no conflicts of interest. References 1. Ding Y, Li Y, Yu G (2016) Exploring bio-inspired quinone-based organic redox flow batteries: a combined experimental and computational study. Chem 1:790–801 2. Young DC (2009) Computational drug design: a guide for computational and medicinal chemists. John Wiley & Sons, Hoboken, NJ 3. Chan WK, Lorenzi PL, Anishkin A, Purwaha P, Rogers DM, Sukharev S, Rempe SB, Weinstein J (2014) The glutaminase activity of L-asparaginase is not required for anticancer activity against ASNS-negative cells. Blood 123:3596–3606 4. Anishkin A, Vanegas JM, Rogers DM, Lorenzi PL, Chan WK, Purwaha P, Weinstein JN,

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39. Phillips JC, Braun R, Wang W, Gumbart J (2005) Scalable molecular dynamics with NAMD. J Comput Chem 26:1781–1802 40. Grossfield A (2008) An implementation of WHAM: the weighted histogram analysis method. http://membrane.urmc.rochester. edu/Software/WHAM/WHAM.html 41. Feller SE, Zhang Y, Brooks BR (1995) Constant pressure molecular dynamics simulation: the Langevin piston method. J Chem Phys 103:4613–4621 42. K€astner J (2011) Umbrella sampling. Wiley Interdiscip Rev Comput Mol 1:932–942 43. Beutler TC, van Gunsteren WF (1994) The computation of a potential of mean force: choice of the biasing potential in the umbrella sampling technique. J Chem Phys 100:1492–1497 44. Humphrey W, Dalke A, Schulten K (1996) VMD—visual molecular dynamics. J Mol Graph 14:33–38

Chapter 3 Nanodisc Reconstitution of Channelrhodopsins Heterologously Expressed in Pichia pastoris for Biophysical Investigations Maria Walter and Ramona Schlesinger Abstract For a successful characterization of channelrhodopsins with biophysical methods like FTIR, Raman, EPR and NMR spectroscopy and X-ray crystallography, large amounts of purified protein are requested. For proteins of eukaryotic origin, which are poorly expressing in bacterial systems or not at all, the yeast Pichia pastoris represents a promising alternative for overexpression. Here we describe the methods for cloning, overexpression and mutagenesis as well as the purification procedures for channelrhodopsin-2 from Chlamydomonas reinhardtii (CrChR2), channelrhodopsin-1 from Chlamydomonas augustae (CaChR1) and the scaffold protein MSP1D1 for reconstitution of the membrane proteins into nanodiscs. Finally, protocols are provided to study CaChR1 by FTIR difference spectroscopy and by time-resolved UV/Vis spectroscopy. Key words Channelrhodopsin, Nanodiscs, Expression, Mutagenesis, Purification, Membrane protein reconstitution, Pichia pastoris, Steady-state FTIR, Time-resolved UV/Vis spectroscopy

1

Introduction Channelrhodopsins (ChRs) are photo-responsive cation channels, which have been originally discovered in the alga Chlamydomonas reinhardtii. ChR is located in the eyespot of the latter where it is involved in phototaxic responses of the cells [1, 2]. Shortly after their discovery, they were identified as being outward directed cation channels with the ability to depolarize cells [3]. Pioneering work expressing channelrhodopsins in different cell types like neurons to provoke physiological responses by light excitation has been first described in 2005 [4, 5] and is subsequently termed “optogenetics”. The first work already pointing to this kind of applications has already been undergone as early as 1994 in the group of Georg Bu¨ldt [6] with the proton pump bacteriorhodopsin in mitochondrial membranes. Nowadays many different channelrhodopsins from various organisms are described with different characteristics [7]. Even

Robert E. Dempski (ed.), Channelrhodopsin: Methods and Protocols, Methods in Molecular Biology, vol. 2191, https://doi.org/10.1007/978-1-0716-0830-2_3, © Springer Science+Business Media, LLC, part of Springer Nature 2021

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anion channnelrhodopsins can be found in nature, which might be used as optogenetic tools for membrane hyperpolarization [8]. Here we will focus on the construction, expression and purification of the most popular and frequently used CrChR2 (channelrhodopsin-2 from Chlamydomonas reinhardtii) and of CaChR1 (channelrhodopsin-1 from Chlamydomonas augustae), the latter exhibiting a red-shifted absorption maximum for improved penetration depth in organismic applications [7]. Both proteins can be expressed well in the heterologous host Pichia pastoris, which is a methylotrophic organism and allows proteins to be induced from an AOX promoter by methanol. As the cell walls of yeast cells are very hard to break, a high-pressure cell disrupter of Constant Systems is used at 2.7 kbar, which cracks most of the cells after three passes. The membranes collected via ultracentrifugation are subsequently solubilized with the detergents DM (n-decyl-β-D-maltopyranoside) or DDM (ndodecyl-β-D-maltopyranoside) and purified via Ni-NTA column ¨ KTA system. on an automated A We developed reconstitution protocols of channelrhodopsins into nanodiscs [9], which mimic the natural lipidic bilayer system. We expressed the scaffold protein MSPID1, a modified protein of the human apolipoprotein A1, in Escherichia coli, and purified it via its His tag using affinity chromatography. The lipid used for reconstitution of the membrane proteins was DMPC, which stabilizes both channelrhodopsins described here. The purification protocol of the protein-filled nanodiscs involves size exclusion chromatography, which separates occupied from empty nanodiscs. Single components and nanodiscs, containing dimers of the proteins, can be stored at
80  C for several months without any loss of functionality.

2

Materials To prepare all solutions use highly pure water by purifying water to a resistance of ca. 18 MΩ at room temperature using a Milli-Q apparatus.

2.1 Construction of Recombinant Channelrhodopsins, Cloning and Mutagenesis

1. Oligonucleotides for CaChR1 and CrChR2: Primer a: 50 -GCGAATTCCATATGGATTATGGAGGC GCCCTGAG-30 (forward flanking primer for CrChR2). Primer b: 50 -GCGGCCGCAAGCTTTCAATGGTGATGGT GATGGTGATGGTGATGGTGGCTAGCGCCAGCCT CGGCCTCGTCCTC-30 (reverse flanking primer for CrChR2). Primer c: 50 -GCCATATGGAATTCACCATGGACACTTT GGCTTGGGTTGCTAG-30 (forward flanking primer for CaChR1).

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Primer d: 50 -GAATTCGCGGCCGCTTAATGATGGTGAT GGTGATGATGGTGGTGATGATCTTCGTC-30 (reverse flanking primer CaChR1). 2. Escherichia coli strain Top 10: genetic markers—F- mcrA Δ(mrr-hsdRMS-mcrBC) Φ80lacZΔM15 Δ lacX74 recA1 araD139 Δ(ara-leu)7697 galU galK rpsL (StrR) endA1 nupG (Thermo Fisher Scientific). 3. Vectors: pGEMHE chop2 full length (kindly provided by Georg Nagel, Wu¨rzburg), pCR-Blunt (Thermo Fisher Scientific); pPIC9K (Life Technologies GmbH); pPIC9K-CaChR1 (with codon optimized gene for P. pastoris, coding for amino acids 1352 and a C-terminal 10X His tag, BioCat GmbH). 4. Phusion Polymerase High-Fidelity PCR Kit. 5. Zero Blunt PCR Cloning Kit (Thermo Fisher Scientific). 6. Plasmid Mini Kit. 7. Restriction endonucleases: EcoRI; NotI; DpnI. 8. Agarose gel electrophoresis: agarose gel electrophoresis chamber; 50 TAE-buffer—2 M Tris, 0.05 M EDTA adjusted to pH 8 with acetic acid; GelRed (Biotium); fast digest green buffer (Thermo Fisher Scientific); 1 kb DNA ladder. 9. Gel Extraction Kit. 10. T4 Ligase Kit. 11. Antarctic Phosphatase Kit. 12. Mini centrifuge for Eppendorf caps. 13. PCR cycler. 14. MicroPulser Electroporation Apparatus (Bio-Rad) with electroporation cuvettes (2 mm). 15. BHI medium: 37 g Brain Heart Infusion powder/L. 16. BHI agar: 52 g Brain Heart Infusion agar powder/L. 17. Kan stock solution: 50 mg/mL kanamycin (sterile filtrated). 2.2 Transformation of Yeast Pichia pastoris and Analysis of Clones

1. Pichia pastoris strain SMD1163: relevant genetic markers—Δ his4Δpep4Δprb1; selectable marker—histidine auxotroph (Life Technologies GmbH, Germany). 2. Restriction endonuclease: SalI. 3. Sorbitol solution: 1 M sorbitol (sterile filtrated). 4. MD (minimal dextrose) agar: 1.34% (m/v) YNB (yeast nitrogen base), 4  10
5% (m/v) biotin, 2% (m/v) glucose, 1.5% agar. 5. YPD (yeast extract peptone dextrose) agar with geneticin (¼G418): 1% (m/v) yeast extract, 2% (m/v) peptone, 2%

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(m/v) glucose, 1.5% agar supplemented with variable geneticin concentrations ranging from 0.1 to 4 mg/mL (see Note 1). 6. Lysis buffer for DNA extraction from Pichia pastoris: 0.2 M lithium acetate, 1% SDS (sodium dodecyl sulphate). 7. Vortex. 8. Thermo mixer. 9. Tip-sonicator. 2.3 Expression of Channelrhodopsins

1. BMGY (Buffered Glycerol-complex Medium): 1% (m/v) yeast extract, 2% (m/v) peptone, 100 mM potassium phosphate pH 7.4, 1.34% (m/v) YNB (yeast nitrogen base), 4  10
5% (m/v), biotin, 1% (v/v) glycerol (see Note 2). 2. BMMY (Buffered Methanol-complex Medium): 1% (m/v) yeast extract, 2% (m/v) peptone, 100 mM potassium phosphate pH 7.4, 1.34% (m/v) YNB, 4  10
5% (m/v), biotin, 0.5% (v/v) methanol (see Note 2). 3. Methanol/all-trans retinal mixture: methanol containing 500 μM all-trans retinal (from a stock solution of 25 mM all-trans retinal in ethanol). 4. Amp stock solution: 200 mg/mL ampicillin (sterile filtrated). 5. Shaker and baffled flasks. 6. Cooling centrifuge for harvesting cells.

2.4 Purification of Channelrhodopsins

1. Breaking buffer: 200 mM sodium phosphate, 1 mM EDTA, 5% glycerol, pH 7.4. 2. cOmplete, EDTA-free protease inhibitor cocktail tablet (Roche Applied Science). 3. PMSF (phenylmethylsulfonyl fluoride) stock solution: 20 mg/ mL PMSF in isopropanol. 4. Benzamidine stock solution: 20 mg/mL benzamidine. 5. Cell Disrupter (TS Series, 1.1 kW; Constant Systems). 6. Solubilization buffer for CaChR1: 20 mM HEPES pH 7.4, 100 mM NaCl, 2% DDM. 7. Solubilization buffer for CrChR2: 20 mM HEPES pH 6.0, 100 mM NaCl, 2 M urea, 1% DM. 8. Dounce homogenizer. 9. 5 mL column with Ni-NTA sepharose. ¨ KTA avant 25 (GE Healthcare). 10. A 11. Wash buffer: 20 mM HEPES, 100 mM NaCl pH 7.4, 0.03% DDM (for CaChR1), 0.2% DM (for CrChR2). 12. Elution buffer: 20 mM HEPES, 100 mM NaCl pH 7.4, 0.03% DDM (for CaChR1), 0.2% DM (for CrChR2) and 500 mM imidazole.

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13. UV/Vis spectrophotometer. 14. Ultracentrifuge. 2.5 Concentration of Purified Channelrhodopsins and Removal of Excess Detergent

1. Centrifugal filter units/concentrators (for different volumes; molecular weight cutoff 50 kDa).

2.6 Expression of Scaffold Protein MSP1D1

1. Escherichia coli strain BL21::DE3 Codon+ RIL: genetic markers—argU (AGA, AGG), ileY (AUA), leuW (CUA); selectable marker: chloramphenicol (Agilent Technologies, Inc., Santa Clara, California, USA).

2. Cooling centrifuge with swing out rotor for 15 and 50 mL tubes. 3. Filter wash buffer: 2 mM HEPES pH 7.4, 5 mM NaCl.

2. Vector: pMSP1D1 (a pET 28a derivative [10]). 3. TB (Terrific Broth) medium: 1.2% (m/v) tryptone, 2.4% (m/v) yeast extract, 0.5% (v/v) glycerol, 89 mM potassium phosphate pH 7.4 (see Note 3). 4. IPTG (Isopropyl-β-D-thiogalactopyranosid) stock solution: 1 M IPTG (sterile filtrated). 2.7 Purification of MSP1D1 and Reconstitution to Nanodiscs

1. MSP buffer: 40 mM Tris–HCl pH 8.0, 300 mM NaCl. 2. MSP/Triton buffer: MSP buffer, 1% Triton-X100. 3. Cannula (0.25 mm). 4. Sterile filters (0.2 μm). 5. Centrifugal filter units (10 kDa and 50 kDa cutoff). 6. MSP/sodium cholate buffer: MSP buffer, 50 mM sodium cholate. 7. MSP/EDTA (Ethylenediaminetetraacetic acid) buffer: MSP buffer, 0.5 mM EDTA. 8. Size exclusion column HiLoad™ Superdex 200 16/60 (GE Healthcare). 9. Size exclusion column HiLoad™ Superdex 200 10/300 GL (GE Healthcare). 10. HEPES buffer: 20 mM HEPES pH 7.4, 100 mM NaCl. 11. Cholate solution: 200 mM sodium cholate in HEPES buffer. 12. Lipid: 50 mM DMPC (1,2-Dimyristoyl-sn-glycero-3-phosphorylcholine) in HEPES buffer and 100 mM sodium cholate. 13. Bio-Beads SM-2 resin (Bio-Rad).

2.8 FTIR Difference Spectroscopy

1. BaF2 windows (20 mm diameter). 2. FTIR spectrometer (Vertex 80v, Bruker). 3. LED emitting at 505 nm.

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Maria Walter and Ramona Schlesinger

1. Flash photolysis setup (LKS80, Applied Photophysics).

2.9 Time-Resolved UV/Vis Spectroscopy

3

2. Nd:YAG laser.

Methods 1. Mix the following components to synthesize recombinant CrChR2 (including only the N-terminal membraneous part containing amino acids 1–307 and a 10 His tag): 1 μL template (pGEMHE chop2 full length) containing cDNA of CrChR2, 1 μL with 100 pmol of primer a, 1 μL with 100 pmol of primer b, 10 μL 5 HF buffer of the PCR kit, 1 μL dNTPs (10 mM each), 5 μL DMSO, 0.5 μL Phusion Hot Start Polymerase and fill up to a total volume of 50 μL with Milli-Q purified water. Run the following programme in the PCR cycler: 1  98  C for 3 min, 45  98  C for 10 s and 72  C for 30 s, finally 1  72  C for 5 min. Primer a incorporates an EcoRI and a NdeI restriction site upstream of the start codon. Primer b adds an additional sequence for C-terminal NheI site (for eventually exchanging tags) and the10 His-tag coding sequence upstream of the stop codon and a HindIII and NotI site downstream of the stop codon (see Fig. 1).

3.1 Construction of Recombinant Channelrhodopsin-2

2. Purify the PCR product on a 1% agarose gel in 1 TAE buffer containing GelRed (as instructed by the manufacturer). 3. Cut out the band containing the PCR product from the gel with a scalpel, and purify the DNA with a Gel Extraction Kit (as instructed by the manufacturer) and elute with 30 μL elution buffer. 4. Ligate 7.5 μL of the PCR product with 0.5 μL of the pCR-Blunt vector (from Zero Blunt PCR cloning Kit) with

EcoRI

NdeI

Stop aa 307

primer 1

aa 737

CrChR2 full length NheI Start

Stop NotI

primer 2

10xHis tag

HindIII

Stop

CrChR2

10xHis tag

Start

Fig. 1 Scheme for the construction of recombinant CrChR2 by PCR

recombinant CrChR2

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35

1 μL ligase and 1 μL 10 ligase buffer from the T4 Ligase Kit for at least 30 min at room temperature. 5. Electro-transform 3–4 μL of the ligation into 50 μL salt-free E. coli Top 10 cells with a MicroPulser. Add 1 mL of BHI medium, and shake for 1 h at 37  C before spreading on BHI-agar plates supplemented with 50 μg/mL kanamycin (from kan stock solution). 6. Select single colonies from the agar plates, and grow them overnight in 3–4 mL of BHI medium supplemented with 50 μg/mL kanamycin. Isolate plasmid DNA with the Plasmid Mini Kit (as instructed by the manufacturer) and elute with 100 μL elution buffer. 7. Analyse plasmids by digestion with EcoRI and subsequent run on an agarose gel. 8. Send plasmids with correct size for sequencing to a company. 9. Use the plasmid with verified recombinant CrChR2 gene as template for generating CrChR2 variants with the megaprimer method and cloning into expression vector. 3.2 Cloning into the Expression Vector

1. Subclone CrChR2 (and variants) from the pCR-Blunt vector (from Subheading 3.1) in the expression vector pPIC9K behind the methanol inducible AOX promotor. For this, cut recombinant CrChR2 sequences from 20 μL of the corresponding pCR-Blunt construct with EcoRI and NotI in a total volume of 40 μL. 2. Digest 20 μL vector pPIC9K (out of a total of 100 μL Mini Plasmid Prep) with 2 μL of EcoRI and 2 μL NotI in a total volume of 40 μL (following the manufacturer’s instruction). To dephosphorylate the 50 ends, add 2 μL Antarctic Phosphatase after approx. 30 min, and incubate at 37  C for additional 30 min. 3. Purify the CrChR2 fragment and the digested, dephosphorylated pPIC9K in correspondence of steps 2 and 3 in Subheading 3.1. 4. Mix the ligation reaction by adding 15 μL CrChR2-DNA (out of 30 μL of the eluted fragment) to 2 μL pPIC9K (out of 30 μL linearized vector) with 2 μL 10 ligase buffer and 1 μL T4 ligase in a total volume of 20 μL. Incubate the ligation mixture at room temperature for at least 30 min (see Note 4). 5. Continue as described under steps 5–8 in Subheading 3.1, except for the digestion of individual plasmids, which need to be done with EcoRI and NotI. 6. Use the plasmid with verified recombinant CrChR2 gene as template for generating CrChR2 variants with the QuikChange method and for expression in yeast cells.

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3.3 Mutagenesis of Channelrhodopsins with the Megaprimer Method

To introduce single mutations for different CrChR2 variants like used in [11, 12] and for CaChR1 variants, use the plasmids with recombinant channelrhodopsins as template. 1. Design mutagenesis primers with the mutation in the middle of the primer flanked by sequences with a calculated melting temperature on each site of approximately 45–50  C (by calculating 2 for T and A and 4 for C and G), and order them from a company (see Note 5). 2. Set up a PCR-1 reaction as described in Subheading 3.1, step 1 with the plasmid containing the recombinant gene and as primers the mutagenesis primer and a flanking primer (listed under Subheading 2.1, item 1) enclosing the fragment which should be amplified first and purify as described under Subheading 3.1, steps 2 and 3. 3. Set up a PCR-2 reaction mainly as described under step 2 but using as primers half of the fragment received from PCR-1 (15 μL) and the other flanking primer of the gene, which was not used in PCR-1, to enclose the whole gene and purify as described under Subheading 3.1, steps 2–8. 4. Use the plasmid with verified mutagenized ChR gene for cloning into expression vector following the protocol described under Subheading 3.2, steps 1–5. 5. Use the verified expression plasmids with mutagenized CaChR1 gene or CrChR2 variants for transformation and expression in yeast cells.

3.4 Mutagenesis of Channelrhodopsins with the QuikChange Method (see Note 6)

1. Mix the following components to mutagenize recombinant CrChR2 or CaChR1 in a QuikChange PCR: 1 μL expression vector containing the gene as template, 1.5 μL with 10 pmol of forward mutagenesis primer, 1.5 μL with 10 pmol of reverse mutagenesis primer, 10 μL 5 HF buffer of the PCR kit, 1 μL dNTPs (10 mM each), 2.5 μL DMSO and 0.5 μL Phusion Hot Start Polymerase and fill up to a total volume of 50 μL with Milli-Q purified water. Run the following programme in the PCR cycler: 1 98  C for 3 min, 18  98  C for 1 min, 60  C for 1 min and 72  C for 5 min, finally 1  72  C for 8 min. 2. Add 1 μL DpnI to the reaction to digest methylated template for at least 1 h and purify following Subheading 3.1, steps 2 and 3. 3. Electro-transform 10 μL of the digested PCR product, and make selection of colonies mainly as described in Subheading 3.1, steps 5 and 6. 4. Analyse plasmids as indicated under Subheading 3.1, step 7 but with NotI and EcoRI and continue as indicated under Subheading 3.1, step 8.

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expression host: Pichia pastoris

verification by sequencing

nucleus

AOX-promotor

PCR pBlunt

1.

pPIC9K

2.

3.

multiple copy inserts

4.

Fig. 2 Workflow from PCR to the expression strain 3.5 Transformation into Yeast Cells

1. Linearize ¼ of a mini-DNA preparation of the expression plasmid with SalI and purify following Subheading 3.1, steps 2 and 3. 2. Electro-transform 10 μL of the linearized plasmid into 50 μL salt-free SDM1163 cells with a MicroPulser, and add immediately 1 mL sorbitol solution after the pulse. 3. Pellet the cells, resuspend them in 200 μL sorbitol solution, plate them on MD agar plates, and let them grow for 2 days at 30  C. 4. Add 5 mL YPD medium (see Note 1) on the MD agar plate, scratch 1/3 of the cell layer, and pipet the suspension into a culture tube (see Note 7). 5. Incubate at 30  C and 150 rpm for 1 h. 6. Plate 100 μL of cell suspension on YPD agar containing 0.1 mg/mL geneticin, and incubate at 30  C. 7. Continue the next days to transfer growing colonies to higher concentrations of geneticin to select hyper-resistant clones with multicopy gene inserts (see Note 8). The schematic workflow from PCR to generate channelrhodopsin constructs up to the hyper-resistant expression hosts is summarized in Fig. 2 (adapted from [13]).

3.6 Extraction of Genomic DNA from Pichia pastoris

The protocol was adapted from [14]. 1. Use 250–400 μL cell suspension in YPD with an OD600 of 0.4–0.6. 2. Disrupt the cells with a tip-sonicator. 3. Centrifuge at 15,000  g for 1 min at room temperature. Discard the supernatant. 4. Resuspend the pellet in 100 μL lysis buffer, and incubate for 5–10 min at 70  C. 5. Add 300 μL 96% ethanol and vortex for 10 s. 6. Centrifuge at 15,000  g for 3 min at room temperature. Discard the supernatant. 7. Resuspend the pellet in 300 μL 70% ethanol.

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Maria Walter and Ramona Schlesinger

8. Centrifuge at 15,000  g for 3 min at room temperature. Discard the supernatant. 9. Dry the pellet. 10. Resuspend the pellet in 30 μL Milli-Q water. 11. Centrifuge at 15,000  g for 15 s at room temperature. 12. Use 1 μL of the supernatant as template for a PCR reaction with flanking primers (listed under Subheading 2.1, item 1). 3.7 Expression of Channelrhodopsins

1. Inoculate 50 mL of YPD medium with the expression strain from an agar plate, and incubate until late logarithmic phase at 30  C and 180 rpm (see Note 9). 2. Dilute the 50 mL YPD culture into 500 mL BMGY medium in a 2 L baffled flask supplemented with 200 μg/mL ampicillin and 50 μg/mL kanamycin (from amp and kan stock solutions), and incubate overnight at 30  C, 180 rpm. Antibiotics are not necessary but advantageous to avoid bacterial contaminations. 3. In the morning, dilute by adding 100 mL BMGY culture into 500 mL BMMY with antibiotics (see step 2) in 2 L baffled flasks, and feed additionally with 2.55 mL methanol/all-trans retinal mixture. Incubate at 30  C, 180 rpm (see Note 10). 4. In the evening add again 2.55 mL methanol/all-trans retinal as under step 3 (second feed). Continue to incubate. 5. Next day feed twice a day (third feed in the morning and fourth feed in the evening) as recommended under step 3. Continue to incubate overnight. 6. Harvest the cells by centrifugation at 6000  g for 10 min. We obtain ca. 15–20 g cells per litre culture. 7. Store the cells at
80  C or continue with protein purification.

3.8 Disruption, Solubilization and Purification on Ni-NTA Sepharose

1. Resuspend ca. 30 g cell pellet (CaChR1) or up to 100 g cell pellet (CrChR2) in 100–300 mL breaking buffer supplemented with 20 μg/mL PMSF (from PMSF stock solution) and 10 μg/mL benzamidine (from benzamidine stock solution) (see Note 11). 2. Pass the suspension 5 through a cell disruptor at a pressure of 2.7 kbar until most of the cells are cracked and add immediately a cOmplete protease inhibitor tablet resolved in 1 mL of water (see Note 12). 3. Remove the cell debris by centrifugation at 5250  g for 10 min, 4  C. Collect the supernatant. 4. Wash the pellet with additional amount of breaking buffer and centrifuge as before. Pool with the supernatant from step 3. 5. Collect the membranes by a second high speed centrifugation step (186,000  g, 3–4 h at 4  C).

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39

6. Determine the weight of the pellets containing the protein membranes, and resuspend in 9 excess of solubilization buffer supplemented with PMSF (as indicated under step 1). 7. Homogenize in a Dounce homogenizer, and stir for at least 1 h up to overnight at 4  C. 8. If you like to freeze at
80  C after membrane collection, then use solubilization buffer without detergent in step 6, and homogenize and add the 2% DDM for CaChR1 or 1% DM for CrChR2 not until the membranes are unfrozen for solubilization as described in step 7. 9. Centrifuge the solubilization mixture for 1 h at 186,000  g (4  C). 10. Apply the supernatant supplemented with imidazole (20 mM for CaChR1, 60 mM for CrChR2) several times onto an ¨ KTA avant equilibrated 5 mL Ni-NTA column in the A 25 with a flow rate of 5 mL/min at room temperature. 11. Execute washing steps for CrChR2 with 5 CV (column volume) wash buffer supplemented with 60 mM imidazole followed by 5 CV of wash buffer supplemented with 120 mM imidazole and for CaChR1 5 CV wash buffer with 20 mM imidazole followed by 5 CV of wash buffer with 50 mM imidazole (flow rate 4 mL/min). 12. Elute the protein with a linear gradient of the last wash buffer and the elution buffer over 50 mL using reverse flow. 13. Pool the samples in a concentrator (50 kDa cutoff), and concentrate by spinning down. 14. Add wash buffer and spin down to remove the imidazole (see Note 13). 15. Analyse the samples by measuring a spectrum on a UV/Vis spectrophotometer in the range of 250–700 nm. 16. Calculate the concentration by using the Lambert-Beer law and specific extinction coefficients (see Note 14) and the purity by calculating the ratio of the absorbance at 280 nm divided by the specific absorbance of the protein. 17. Additionally analyse the samples for purity on SDS-PAGE (see Fig. 3 for CaChR1 and Note 15). 18. Apply size exclusion chromatography to achieve higher optical purity when needed. 19. Freeze in liquid nitrogen and store at
80  C. 3.9 Concentration of the Proteins and Removal of Excess Detergent for FTIR Spectroscopy

1. Use centrifugal filter units to concentrate channelrhodopsins. 2. Centrifuge at 5000  g for 5 min at 4  C. Remove the filtrate, mix the protein solution gently with a pipette tip, add 5 excess (v/v) of filter wash buffer, and repeat this step five times.

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Maria Walter and Ramona Schlesinger kDa

1

2

180 135 100 75

dimeric CaChR1

63 48

monomeric CaChR1

35

25

Fig. 3 SDS-PAGE of recombinant CaChR1 after affinity purification on Ni-NTA sepharose column. Lane 1 shows a molecular weight standard (in kDa) and lane 2 the purified CaChR1

3. Determine the reduced detergent content with ATR spectroscopy (see Note 16). Detergent content 2% is preferred. 4. Proceed with the IR transmission experiments or aliquot the protein, freeze in liquid nitrogen, and store at
80  C. 3.10 Expression of Scaffold Protein MSP1D1

1. Electro-transform 1 μL pMSP1D1 into 50 μL salt-free E. coli strain BL21::DE3 Codon+ RIL with a MicroPulser. Add 1 mL of BHI medium, and shake for 1 h at 37  C. 2. Spread the cells on a large BHI-agar plate supplemented with 50 μg/mL kanamycin, and incubate overnight at 37  C. 3. Scratch the cell layer and resuspend in 40 mL TB medium. 4. Inoculate 500 mL TB medium supplemented with 50 μg/mL kanamycin with 10 mL of the cell suspension in a 2 L flask, and incubate at 37  C, 150 rpm. 5. When OD600 of 0.8–1 is reached, induce the MSP1D1 expression with 1 mM IPTG (from IPTG stock solution). 6. Incubate the culture for another 4 h at 37  C, 150 rpm. 7. Harvest the cells via centrifugation at 8000  g, 4  C, 15 min. 8. Store cell pellets at
80  C.

3.11 Purification of MSP1D1

1. Thaw cells in MSP/Triton buffer and lyse by sonication. Use an ice bath to prevent overheating of the suspension. 2. Remove the cell debris by centrifugation at 125,000  g, 4  C, 20 min.

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41

3. Add 20 μg/mL (end concentration) PMSF and 10 μg/mL (end concentration) benzamidine to the supernatant. 4. Filter the supernatant particle free, and apply to Ni-NTA affin¨ KTA avant 25 system. ity chromatography on A 5. Load the supernatant three times on a 5 mL Ni-NTA sepharose column (flow rate 4 mL/min) equilibrated with MSP buffer. 6. Wash the column with four CV of each: MSP/sodium cholate buffer, MSP buffer and MSP buffer containing 50 mM imidazole (flow rate 5 mL/min). 7. Elute MSP1D1 applying a linear gradient of 50–300 mM imidazole in MSP buffer (flow rate 4 mL/min). 8. Concentrate the protein in a centrifugal filter unit to reduce the volume. 9. Load MSP1D1 on the size exclusion column Superdex 200 16/60 previously equilibrated with MSP/EDTA buffer (flow rate 1 mL/min). 10. Pool MSP1D1 containing fractions and concentrate in a centrifugal filter unit. 11. Measure the absorbance at 280 nm and calculate the concentration using the extinction coefficient at 280 nm of 21,000 M
1 cm
1. 12. Adjust the concentration to 200 μM and aliquot. Freeze aliquots in liquid nitrogen and store at
80  C. 3.12 Reconstitution of CaChR1 or CrChR2 in MSP1D1 DMPC Nanodiscs

The protocol is adapted from [15, 16]. 1. Mix CaChR1 (~200 μM) or CrChR2 (~200 μM), MSP1D1 (200 μM) and lipid DMPC in the following molar ratio: CaChR1:MSP1D1:DMPC 1:1:50 (for CaChR1 nanodiscs); CrChR2:MSP1D1:DMPC 0.5:1:55 (for CrChR2 nanodiscs); and MSP1D1:DMPC 1:80 (for empty nanodiscs). 2. Add cholate solution to adjust the concentration of cholate to 20 mM. 3. Incubate the protein/lipid solution for 1 h at 25  C, 500 rpm in a thermo mixer. 4. Add 1 g Bio-Beads (equilibrated with HEPES buffer) per mL protein/lipid solution, and incubate for 2 h at 25  C, 500 rpm. During this step the assembly of the nanodiscs takes place. 5. Transfer the nanodiscs solution from the Bio-Beads into an Eppendorf cap using a cannula. Rinse the Bio-Beads with HEPES buffer, and combine this wash with the first solution in the Eppendorf cap.

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Maria Walter and Ramona Schlesinger

Fig. 4 Size exclusion chromatography profile of nanodiscs containing CaChR1 (red), CrChR2 (blue) as well as empty nanodiscs (grey)

6. Reduce the volume by centrifugation in a concentrator (50 kDa cutoff) if needed. 7. Centrifuge for 15 min at 15,000  g, 4  C to remove larger aggregates. 8. Apply to the size exclusion column Superdex 10/300 GL previously equilibrated with HEPES buffer. 9. Monitor the absorbance at 280 nm. Typical profiles of channelrhodopsin containing nanodiscs are depicted in Fig. 4. 3.13 FTIR Difference Spectroscopy

See [17] for the procedure of an IR transmission experiment. 1. Pipet 2–5 μL of detergent solubilized and washed (Subheading 3.6, step 2) protein on the BaF2 window. 2. Dry the protein film under gentle air flow. 3. Rehydrate the protein film with 3–4 μL of glycerol/water (2/8 w/w) mixture distributed in droplets near the film without contact to it. 4. Place a thin silicone O-ring (~1 mm thickness) on top of the BaF2 window and seal the sample with a second BaF2 window. 5. Place the sandwich sample into a holder. Place the holder into the sample compartment of a FTIR spectrometer. 6. Wait 30 min to 1 h until the hydration level of the protein film stabilizes before performing light-induced FTIR experiments.

Production of ChRs, Nanodisc Reconstitution and Spectroscopy

43

Fig. 5 Light-induced FTIR difference absorbance spectrum of detergent-solubilized CaChR1. Negative bands represent the vibrations of the dark state; positive bands represent the vibration of the photo intermediate. Dashed fields highlight the important vibrations in CaChR1

7. Repeat the following illumination protocol to a final of 3000 co-added scans: keep the sample in the dark for 10 s, record the spectra in the last 5 s, illuminate subsequently for 10 s with an LED emitting at 505 nm, and record the spectra in the last 5 s. 8. Calculate the difference absorbance ΔA according to the following equation ΔA ¼
log

I light I dark

where Idark and Ilight are the transmitted IR intensities in the dark state and a photo intermediate, respectively (see Fig. 5 for a FTIR difference spectrum of CaChR1 over the complete mid-infrared region [18]). 3.14 Time-Resolved UV/Vis Spectroscopy

1. Place a cuvette with detergent solubilized CaChR1 (in wash buffer, see Subheading 2.4.11) in a sample compartment of a flash photolysis setup. 2. Excite the sample with a 10 ns laser pulse of a Nd:YAG laser. Adjust the laser pulse to 532 nm with an optical parametric oscillator (OPO). Set the energy density of the laser pulse to 3 mJ/cm2. 3. Operate the xenon arc lamp in a pulsed mode for the early time scales (100 ns–1 ms) and in the continuous mode for the later time scales (1 ms–10 s).

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Maria Walter and Ramona Schlesinger

Fig. 6 Light-induced UV/Vis absorbance changes of detergent-solubilized CaChR1 from 100 ns to 5 s. Shown are the ground state depletion (CaChR1520), the early red-shifted photo intermediate (P1590) and the blueshifted photo intermediate (P2380)

4. Measure the kinetics for each wavelength before and after excitation, and record the absorbance difference (light minus dark). 5. Record 20,000,000 data points and average logarithmically for each kinetic. Average each kinetic 5–10 times to reduce the signal-to-noise ratio. 6. Merge the recorded kinetics for every wavelength (early and late time scales; see Fig. 6).

4

Notes 1. Only after the medium (yeast extract, peptone and agar) has been autoclaved, sterile-filtered glucose (20% w/v stock solution) should be added. For geneticin addition you should wait until the medium mixture has cooled down to ca. 60  C. When preparing agar plates, be careful with mixing the components to not producing too much bubbles. Correspondingly, YPD medium is prepared as described but without agar and geneticin. 2. The components need to be sterilized separately. Autoclaved peptone together with yeast extract is mixed with separately autoclaved potassium phosphate, YNB and glycerol

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45

(in BMGY). Methanol (in BMMY) can be added in sterilized water. Supplemented biotin needs to be sterile-filtrated. 3. Dissolve tryptone, yeast and glycerol in 900 mL of water and autoclave. 100 mL potassium phosphate pH 7.4 is prepared by dissolving 2.31 g KH2PO4 and 12.54 g K2HPO4 in water and autoclaved separately. Mix both solutions to give 1 L of buffer. 4. To test the quality of the vector fragment, a ligation control by adding all components but replacing the insert with deionized water should be performed. 5. It is up to you which mutagenesis primer you order the forward primer representing the upper DNA strand or the reverse primer representing the lower strand. Usually it is more favourable to produce the larger gene fragment first. You can also order both mutagenesis primers complementary to each other if you also would like to try the QuikChange method for mutagenesis. 6. Once you have your gene in the expression vector, this method is much faster than the megaprimer method described under Subheading 3.3. But depending of the mutagenesis position and sequence, it might not lead to 100% of clones with the desired mutation but to a variable amount of clones with the original sequences given in the template. 7. Alternatively to the method where a pool of cells is transferred on geneticin plates, single colonies can individually be transferred which sometimes gives better results. But be aware that only 1–10% of transformants are hyper-resistant due to multicopy insertion. 8. Try to continue selection up to 4 mg/mL geneticin on agar plates. 9. This pre-preculture is enough for approx. 2.5 L of main culture. For higher volumina plan accordingly more of the starting culture. 10. The expression of the culture in YPD is repressed due to glucose. The cells experience derepression in BMGY because of lack of glucose. Induction of expression takes place only in BMMY as the AOX promoter in front of the gene is methanol responsive. If cultures are foaming too much, you can add 200 μL antifoam to each flask. 11. The used cell mass is depending on the expression rate which is much better for CaChR1 than CrChR2. This step needs adaptation for each variant as the used column volume is typically 5 mL bed volume, and the expressed protein should neither exceed the capacity of the column nor go too much below as

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Maria Walter and Ramona Schlesinger

the latter case leads to impure protein. Of course you can choose bigger or smaller columns to adapt. 12. To observe whether disruption was enough efficient, you can check the lysed cells under the microscope. The unbroken cells are round and can be spotted easily in the partially lysed suspension. Do not add the protease inhibitor cocktail before disruption as this leads to foaming during disruption which makes the procedure less efficient. 13. One washing step with imidazole-free buffer is sufficient, if you proceed with size exclusion chromatography contemporary. Otherwise repeat the washing step several times, and store the protein at
80  C. Another method to remove imidazole is the use of a desalting column, e.g. 10DG column (Bio-Rad) according to manufacturer’s instructions. 14. Values for calculation of concentrations: CaChR1: Extinction coefficient at 36,000 M
1 cm
1; m ¼ 40,660 g/mol.

518

nm—

CrChR2: Extinction coefficient at 470 nm—45,000 M
1 cm
1; m ¼ 35,675 g/mol. 15. For SDS-PAGE analysis, add 5 μL of 5 SDS sample buffer to 20 μL of channelrhodopsin samples, and mix thoroughly. Load onto 12% SDS-PAGE gel using the standard protocol, stain with Coomassie blue, and visualize the protein bands after destaining.

Fig. 7 FTIR difference spectra of DDM solutions (0.1–10% DDM in 2 mM HEPES pH 7.4, 5 mM NaCl) after subtraction of the spectrum of DDM free buffer (left). Correlation of the band intensity at 1030 cm
1 with DDM concentration (right)

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47

16. Use ATR accessory (here Ge-ATR cell) in a FTIR spectrometer (here Vertex 70, Bruker) to measure the intensity of the 1030 cm
1 band in solutions with different detergent concentrations. See [17] for the handling of the ATR cell. First, record a spectrum of 2 μL detergent-free buffer (2 mM HEPES pH 7.4, 5 mM NaCl) in the region 4000–1000 cm
1. Next, use 2 μL of DDM solutions (0.1–10% DDM in 5 mM HEPES pH 7.4, 2 mM NaCl) to record the spectra (4000–1000 cm
1). Scale all spectra to the band at 3353 cm
1 to eliminate the impact of water. Subtract the spectrum of detergent-free buffer from the spectrum of each DDM solution (Fig. 7). The intensity of the band at 1030 cm
1 is proportional to the DDM concentration. Next, record a spectrum of your protein sample with reduced detergent content. Subtract the spectrum of detergent-free buffer from the spectrum of your protein sample. Determine the detergent content in your protein sample using the intensity of 1030 cm
1 band.

Acknowledgements We thank Nils Krause and Vera Muders for developing protocols and Kirsten Hoffmann and Dorothea Heinrich for technical assistance. Financial support was granted from the Deutsche Forschungsgemeinschaft (SFB1078 TP B4). References 1. Sineshchekov OA, Jung KH, Spudich JL (2002) Two rhodopsins mediate phototaxis to low- and high-intensity light in Chlamydomonas reinhardtii. Proc Natl Acad Sci U S A 99:8689–8694 2. Nagel G, Ollig D, Fuhrmann M, Kateriya S, Musti AM, Bamberg E, Hegemann P (2002) Channelrhodopsin-1: a light-gated proton channel in green algae. Science 296:2395–2398 3. Nagel G, Szellas T, Huhn W, Kateriya S, Adeishvili N, Berthold P, Ollig D, Hegemann P, Bamberg E (2003) Channelrhodopsin-2, a directly light-gated cation-selective membrane channel. Proc Natl Acad Sci U S A 100:13940–13945 4. Nagel G, Brauner M, Liewald JF, Adeishvili N, Bamberg E, Gottschalk A (2005) Light activation of channelrhodopsin-2 in excitable cells of Caenorhabditis elegans triggers rapid behavioral responses. Curr Biol 15:2279–2284

5. Boyden ES, Zhang F, Bamberg E, Nagel G, Deisseroth K (2005) Millisecond-timescale, genetically targeted optical control of neural activity. Nat Neurosci 8:1263–1268 6. Hoffmann A, Hildebrandt V, Heberle J, Buldt G (1994) Photoactive mitochondria: in vivo transfer of a light-driven proton pump into the inner mitochondrial membrane of Schizosaccharomyces pombe. Proc Natl Acad Sci U S A 91:9367–9371 7. Hou SY, Govorunova EG, Ntefidou M, Lane CE, Spudich EN, Sineshchekov OA, Spudich JL (2012) Diversity of Chlamydomonas channelrhodopsins. Photochem Photobiol 88:119–128 8. Govorunova EG, Sineshchekov OA, Janz R, Liu X, Spudich JL (2015) NEUROSCIENCE. Natural light-gated anion channels: a family of microbial rhodopsins for advanced optogenetics. Science 349:647–650 9. Nath A, Atkins WM, Sligar SG (2007) Applications of phospholipid bilayer nanodiscs in the

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study of membranes and membrane proteins. Biochemistry 46:2059–2069 10. Denisov IG, Grinkova YV, Lazarides AA, Sligar SG (2004) Directed self-assembly of monodisperse phospholipid bilayer nanodiscs with controlled size. J Am Chem Soc 126:3477–3487 11. Lorenz-Fonfria VA, Resler T, Krause N, Nack M, Gossing M, Fischer von Mollard G, Bamann C, Bamberg E, Schlesinger R, Heberle J (2013) Transient protonation changes in channelrhodopsin-2 and their relevance to channel gating. Proc Natl Acad Sci U S A 110:E1273–E1281 12. Krause N, Engelhard C, Heberle J, Schlesinger R, Bittl R (2013) Structural differences between the closed and open states of channelrhodopsin-2 as observed by EPR spectroscopy. FEBS Lett 587:3309–3313 13. Schlesinger R, Cousin A, Granzin J, BatraSafferling R (2017) Expression and purification of arrestin in yeast Saccharomyces cerevisiae. Methods Cell Biol 142:159–172 ˜ oke M, Kristjuhan K, Kristjuhan A (2011) 14. Lo Extraction of genomic DNA from yeasts for

PCR-based applications. BioTechniques 50:325–328 15. Bayburt TH, Sligar SG (2002) Single-molecule height measurements on microsomal cytochrome P450 in nanometer-scale phospholipid bilayer disks. Proc Natl Acad Sci U S A 99:6725–6730 16. Bayburt TH, Sligar SG (2003) Self-assembly of single integral membrane proteins into soluble nanoscale phospholipid bilayers. Protein Sci 12:2476–2481 17. Lo´renz-Fonfrı´a VA, Heberle J (2014) Proton transfer and protein conformation dynamics in photosensitive proteins by time-resolved stepscan Fourier-transform infrared spectroscopy. J Vis Exp 88:e51622 18. Lorenz-Fonfria VA, Muders V, Schlesinger R, Heberle J (2014) Changes in the hydrogenbonding strength of internal water molecules and cysteine residues in the conductive state of channelrhodopsin-1. J Chem Phys 141:22D507

Chapter 4 Characterizing Channelrhodopsin Channel Properties Via Two-Electrode Voltage Clamp and Kinetic Modeling Lindsey Prignano, Lauren Herchenroder, and Robert E. Dempski Abstract Two-electrode voltage clamp (TEVC) is a preferred electrophysiological technique used to study gating kinetics and ion selectivity of light-activated channelrhodopsins (ChRs). The method uses two intracellular microelectrodes to hold, or clamp, the membrane potential at a specific value and measure the flow of ions across the plasma membrane. Here, we describe the use of TEVC and a simple solution exchange protocol to measure cation selectivity and analyze gating kinetics of the C1C2 chimera expressed in Xenopus laevis oocytes. Detailed instructions on how to process the collected data and interpret the results are also provided. Key words Voltage clamp, Electrophysiology, Ion selectivity, Reversal potential, BAPTA-AM, Kinetic modeling

1

Introduction Electrophysiological techniques allow scientists to measure ion selectivity and gating kinetics of both naturally occurring and synthetic channel forming peptides [1–3]. When studying channelrhodopsins, the two-electrode voltage clamp (TEVC) method is preferred over other common methods such as patch clamp or single-electrode voltage clamp [4, 5]. TEVC measures whole-cell currents across the plasma membrane which is ideal for proteins like channelrhodopsin where the generated currents are too small for single-channel recording [6]. ChR’s relatively small current also enables the use of very large cells required for TEVC, such as Xenopus laevis oocytes, where a high number of ChR channels can populate the plasma membrane. This eliminates the need for expensive fine micromanipulators and simplifies the experimental setup. TEVC uses two intracellular microelectrodes, a potential electrode and a current electrode, that work together to keep the membrane potential fixed at a value specified by the user

Robert E. Dempski (ed.), Channelrhodopsin: Methods and Protocols, Methods in Molecular Biology, vol. 2191, https://doi.org/10.1007/978-1-0716-0830-2_4, © Springer Science+Business Media, LLC, part of Springer Nature 2021

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Fig. 1 Schematic illustration of two-electrode voltage clamp applied to an X. laevis oocyte (left) and a typical photocurrent output of the channelrhodopsin chimera, C1C2, measured at Vm ¼ 100 mV (right). The blue bar indicates channel activation via irradiation with a 470 nm LED light

[2]. Figure 1 illustrates a typical configuration of a TEVC experimental setup using an X. laevis oocyte as an exogenous expression system. The oocyte is submerged in one or more bath solutions (not shown) of varying ionic composition. To clamp the membrane, the membrane potential (Vm) is monitored by impaling the oocyte with the tip of the potential electrode which is connected to the input of a high input impedance voltage follower amplifier (A1). The output of A1 (Vm) is fed to one of the input terminals of the clamping amplifier (A2). A computer connected to a signal generator sends the command voltage signal (Vc) to the other input terminal of A2 according to a protocol defined by the user. A2 is a high-gain differential amplifier which compares Vm to Vc from the two inputs and sends a current proportional to the difference (ε) through the amplifier output. The output current from A2 is then fed through the current electrode which injects the current into the membrane such that Vm matches Vc extremely closely. The amplitude of the current injected into the oocyte to maintain a fixed membrane potential is equal to the current flow across the cell membrane through open ion channels. Therefore, sampling the current signal just upstream of the current electrode gives a timeresolved measure of the net flow of ions through open ChR channels to produce photocurrent traces similar to the one shown on the right-hand side of Fig. 1. The sign of the measured current depends on the charge and direction of the flow of ions across the membrane. Here, we follow the generally accepted convention that the influx of positively charged ions or efflux of negatively charged ions from the cell lends a negatively polarized current. Similarly, the influx of negatively charged ions or efflux of positively charged ions from the cell lends a measured current with a positive polarity.

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Oocytes of the African Clawed frog, Xenopus laevis, are commonly used as an exogenous expression system to study integral membrane proteins and ion channels via electrophysiological techniques [1]. These oocytes are about 1 mm in diameter, large enough to accommodate multiple electrodes required for TEVC and still achieve a voltage clamp at a sufficient rate to study the gating kinetics of the light-activated channel [7]. In addition, X. laevis oocytes have no endogenous light-activated ion channels and therefore provide an ideal environment for recording photocurrents through only channelrhodopsin. This allows the application of a simple ionic solution exchange protocol described herein to easily evaluate the selectivity of ChRs to most mono- and divalent cations using measured reversal potentials. Recording calcium currents, however, poses a unique challenge that requires the addition of a few extra steps in the protocol. X. laevis oocytes express endogenous calcium-activated chloride channels at their plasma membranes [8, 9]. When studying the ion selectivity of ChRs permeable to calcium, the influx of calcium ions from the external solution into the oocyte through open ChR channels can trigger the activation of these endogenous chloride channels [10]. This generates a large outward-rectifying chloride current that can obscure the photocurrent recorded through channelrhodopsin and affect reversal potential measurements. Thus, the contaminating chloride current renders the data invalid for ion selectivity and kinetic analysis of the target ion channel. To work around this issue, a common method is to use an intracellular calcium chelator such as 1,2-bis (2-aminophenoxy)ethane-N,N,N 0 ,N 0 -tetraacetic acid (BAPTA) [1]. BAPTA selectively and efficiently binds Ca2+ in order to buffer the intracellular calcium concentration and thus prevent contaminating currents from the activation of endogenous calciumactivated chloride channels. Since BAPTA cannot penetrate the cell membrane, the BAPTA solution is typically applied via direct injection into the cytosol a few hours prior to TEVC experiments where external solutions with high calcium concentrations will be used. However, repeatedly damaging the membrane with multiple injections poses an added risk to cell health and membrane integrity, which are critical to the success of the experiment. Here, we avoid causing any unnecessary damage to the cell membrane by using the cell-permeant acetoxymethyl ester derivative, BAPTAAM. BAPTA-AM has been demonstrated to effectively buffer intracellular calcium and suppress chloride currents in X. laevis oocytes by other groups in the past [11, 12]. Unlike the parent compound, the acetoxymethyl ester derivative is an uncharged molecule which can diffuse across the plasma membrane. Once inside the cell, the additional groups are cleaved off by endogenous non-specific esterase activity to give the BAPTA free acid form that can bind calcium. This allows BAPTA to be loaded into cells by simply adding BAPTA-AM to the (calcium-free) oocyte medium at least 1 h prior to TEVC experiments.

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Herein, we describe detailed protocols for measuring isolated photocurrents through channelrhodopsin expressed in X. laevis oocytes via two-electrode voltage clamp. Instructions on how to process the recorded data and interpret results are also included. By following the steps below, a wealth of information on gating kinetics and ion selectivity of natural or synthetic channelrhodopsin constructs can be obtained.

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Materials Prepare all solutions with ultrapure water with a resistivity of >18 MΩ∙cm at 25  C.

2.1 In Vitro Synthesis of mRNA for Oocyte Injection

1. Template DNA; 7–10 μg of plasmid DNA containing the gene of interest dissolved in up to 50 μL diethylpyrocarbonate (DEPC)-treated water. Here, we subcloned residues 1-356 of the C1C2 gene and a C-terminal eYFP (Addgene plasmid # 35519) into the vector pTLN using restriction enzymes XbaI and BglII (New England BioLabs). The pTLN vector contains an SP6 RNA polymerase promotor and the MluI restriction enzyme can be used to linearize the plasmid prior to mRNA synthesis as long as the restriction site is not present in the gene of interest. 2. High Pure PCR Product Purification Kit (Roche Life Science). 3. AmpliCap™ SP6 (CELLSCRIPT).

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4. Lithium chloride precipitation solution; 7.5 M lithium chloride, 50 mM EDTA. 5. Agarose. 6. Tris-acetate-EDTA (TAE) running buffer; 40 mM Tris base, 20 mM acetic acid, and 1 mM EDTA, adjusted to pH 8.5. 7. 1% ethidium bromide solution; 10 mg/mL ethidium bromide in water, stored at 4  C away from light. 8. HyperLadderI (BIOLINE) or other suitable DNA ladder and loading dye for gel electrophoresis. 9. Cell-Porator (Life Technologies). 2.2 Preparation of Oocytes and mRNA Injection

1. Oocyte Ringer’s isotonic buffer without calcium (ORI-); 90 mM NaCl, 2 mM KCl, 5 mM MOPS, pH 7.4. 2. Oocyte Ringer’s isotonic buffer with calcium (ORI+); 2 mM CaCl2, 90 mM NaCl, 2 mM KCl, 5 mM MOPS, pH 7.4. 3. Xenopus laevis oocytes freshly harvested from healthy, mature X. laevis female frogs via partial ovariectomy and stored in ORI buffer.

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4. Dissection microscope. 5. Nanoject Injector (Drummond Scientific). 6. Collagenase Type II (Worthington Biochemical) (see Note 1). 7. 100 mg/mL gentamycin stock 8. 1 mM all-trans retinal stock in DMSO 9. Purified 1 ng/nL mRNA in DEPC-treated water. 2.3 Electrophysiology Equipment

1. Dissection microscope. 2. Turbo-tec-03X amplifier (NPI Instruments). 3. Axioscope 1440A digitizer (Axon Instruments). 4. pCLAMP Software (Axon Instruments). 5. LEDMOD V2 300 mW 470  5 nm LED light source guided by a 2 mm fiber optic cable (Omicron). 6. RC-10 bath perfusion chamber (Warner Instruments). 7. 8-channel gravity fed perfusion system with valve controller (Warner Instruments).

2.4 Two-Electrode Voltage Clamp

1. Sodium pH 7.0 bath solution; 115 mM NaCl, 2 mM BaCl2, 1 mM MgCl2, 10 mM HEPES. Adjust to pH 7.0 using 6 M HCl/NaOH. 2. Sodium pH 9.0 bath solution; 115 mM NaCl, 2 mM BaCl2, 1 mM MgCl2, 10 mM Tris. Adjust to pH 9.0 using 6 M HCl/NaOH. 3. Potassium bath solution; 115 mM KCl, 2 mM BaCl2, 1 mM MgCl2, 10 mM HEPES. Adjust to pH 7.0 using 6 M HCl/KOH. 4. Calcium bath solution; 76.7 mM CaCl2, 2 mM BaCl2, 1 mM MgCl2, 10 mM HEPES. Adjust to pH 7.0 using 6 M HCl/KOH. 5. 30 mM BAPTA-AM stock in DMSO. 6. Two glass 1% agar bridges filled with 3 M KCl. 7. 3 M KCl electrode filling solution. 8. Ag/AgCl wire. 9. Clorox Bleach. 10. Borosilicate glass capillaries. 11. PC-12 pipette puller (Narishige).

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1. Prior to mRNA synthesis, the template plasmid DNA containing the gene of interest must be linearized with the appropriate restriction enzyme. Here, we use the MluI restriction site

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present on the pTLN vector for this purpose. In a 1.5 mL Eppendorf tube, digest 10 μg purified plasmid dissolved in 57 μL of reaction buffer with 3 μL MluI or other suitable restriction enzymes. Incubate in a 37  C water bath for 3 h. 2. Confirm successful linearization of the plasmid by gel electrophoresis. Mix 0.3 g agarose with 30 mL TAE running buffer in a small Erlenmeyer flask to make a 1% gel. Heat or microwave the mixture on high for 1.5–2 min or until all the agarose is dissolved. Let stand for 5 min or until cool enough to be handled. Add 30 μL 1% ethidium bromide and swirl to mix. Pour the solution into the gel tray and insert the comb to form wells. Allow the gel to set for at least 30 min. Once the gel has solidified, remove the comb and place the gel tray in the electrophoresis chamber. Fill the chamber with enough TAE running buffer to completely cover the gel. Load 5 μL of HyperLadder I or other appropriate DNA ladder into one of the wells. Load samples of uncut plasmid and the linearization reaction mixture into adjacent wells. Turn on the power and run the gel for 30–45 min or until bands are well separated. Visualize the bands under UV light. Circular, uncut plasmid DNA should travel slightly farther through the gel than linear DNA. 3. Obtain new boxes of freshly autoclaved pipette tips and Eppendorf tubes to use for the remaining procedures. Spray pipettes, benchtop, exterior of tip boxes, and all other materials with 70% ethanol to ensure an RNase-free working environment. 4. Purify the linearized DNA using the Roche High Pure PCR Product Purification Kit. Add 40 μL DEPC-treated water to the digest to reach a final volume of 100 μL, and then add 500 μL binding buffer from the kit. Load the reaction mixture onto a Roche spin column and insert the column into a collection tube. Centrifuge at 13,000  g for 1 min. Discard the flow-through. Add 500 μL wash buffer and centrifuge at 13,000  g for 1 min. Discard the flow-through. Add 200 μL wash buffer and centrifuge at 13,000  g for 1 min. Discard the flow-through. Centrifuge at 13,000  g for an additional 2 min to dry the spin column. Discard the flow-through and insert the column into a new collection tube. Add 50 μL DEPCtreated water to elute the DNA. Let stand for 1 min and then centrifuge at 13,000  g for 1 min. Check the concentration of the purified DNA on a NanoDrop spectrophotometer and concentrate to 300–350 ng/μL in a SpeedVac centrifuge, if needed. 5. Synthesize mRNA from 1 μg of the linearized template DNA using the AmpliCap™ SP6 High Yield Message Maker Kit. Set up the transcription reaction according to kit instructions. Incubate in a 37  C water bath for 2 h.

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6. Add 1 μL RNase-free DNase I from the kit to the reaction mixture and incubate in a 37  C water bath for 15 min. This removes the template DNA from the reaction mixture. 7. Purify the synthesized RNA. Add 30 μL DEPC-treated water and 30 μL lithium chloride precipitation solution to the reaction mixture, and mix gently by pipetting up and down. Incubate at 20  C overnight. Centrifuge at 13,000  g for 30 min in a microcentrifuge chilled to 4  C. There should be a very small white pellet visible at the bottom of the tube. Taking care not to disturb the pellet, remove the supernatant via pipette. Add 150 μL ice-cold 70% ethanol in DEPC-treated water. Centrifuge at 13,000  g for 15 min. Again, carefully remove the supernatant via pipette. Evaporate off any residual ethanol using a SpeedVac at room temperature. Dissolve the RNA pellet in 15 μL DEPC-treated water allowing 10–15 min to fully dissolve. 8. Check the quality of the purified mRNA by running a sample on a 1% agarose gel (see step 2 for gel electrophoresis procedure). RNA of good quality and purity will appear as well-defined bands, while degraded, poor-quality RNA will present as wide streaks through the gel. Quantify the mRNA yield by checking the concentration on a NanoDrop spectrophotometer. 3.2 Preparation of X. laevis Oocytes and mRNA Injection

1. To prepare oocytes for mRNA injection and electrophysiology experiments, the protective outer follicular cell layer and connective tissue surrounding the oocytes must be removed by treatment with Collagenase Type II (see Note 1). In a 150 mm Petri dish, rinse freshly collected oocytes in calcium-free ORIbuffer. Cut the ovarian lobe into small pieces with fine scissors to aid digestion. 2. In an Eppendorf tube, add 100 mg of Collagenase Type II and 1 mL ORI- buffer. Vortex to mix. Filter the mixture through a 0.2 μm sterile syringe filter, and then add the collagenase to the oocyte media for a final concentration of about 3 mg/mL. Incubate with gentle shaking at 17  C for 2–2.5 h. 3. Once the oocytes have fully separated from the connective tissue, rinse the oocytes thoroughly with ORI+ buffer until the solution runs clear to remove all residual collagenase and debris. Incubate oocytes at 17  C in ORI+ solution supplemented with 1 mg/mL gentamycin in the dark overnight (see Note 2). 4. Select 20–30 healthy stage V–VI oocytes for mRNA injection (see Note 3). Inject 50 nL of 1 ng/nL channelrhodopsin mRNA in DEPC-treated water into each oocyte, as shown in Fig. 2. Incubate at 17  C in ORI+ buffer supplemented with 1.5 μM all-trans retinal and 1 mg/mL gentamycin for 48–72 h in the dark to allow for adequate protein expression at the membrane (see Note 4).

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Fig. 2 Healthy stage V oocytes viewed under a dissecting microscope during mRNA injection

5. On the day of electrophysiology experiments, transfer 10–15 injected oocytes to a 35 mm culture dish filled with 5 mL of ORI- buffer while taking care to minimize light exposure. Add 50 μL gentamycin to oocyte media from 100 mg/mL stock (final concentration ¼ 1 mg/mL). Add 16.5 μL of 30 mM BAPTA-AM stock in DMSO (final concentration ¼ 100 μM) to oocyte media and incubate at 17  C for 1–2 h in the dark prior to TEVC experiments. 3.3 Photocurrent Recording Via Two-Electrode Voltage Clamp

1. Deposit a silver chloride coating on the Ag/AgCl wire electrodes by submerging one end of the wires in Clorox bleach for 30 min to 1 h prior to each day of measurements (see Note 5). Rinse the wires with DI water and allow to completely air-dry before use. 2. Fabricate and assemble the current and potential microelectrodes. Using a PC-12 micropipette puller (Narishige), pull microelectrodes from borosilicate glass capillaries. Backfill the electrodes with 3 M KCl and gently tap to remove any air bubbles. Install the electrode onto the holder such that the coated end of the Ag/AgCl wire is submerged in KCl solution and rests near the tip of the electrode. 3. Set up the RC-10 perfusion chamber and check for leaks. 4. Fill the chamber with sodium solution at pH 7.0. Lower the tips of the microelectrodes into the bath solution, and ensure the tip resistance of each electrode is between 0.5 and 1.5 MΩ. If it is not, remove the glass microelectrode and replace it with a new one. Check the tip resistances again to ensure they are within the acceptable range.

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5. Retrieve an oocyte from the dish that has been treated with BAPTA-AM. Place the oocyte in the center of the perfusion chamber, and rinse with ~5–10 mL sodium bath solution before clamping. Position the tips of the two electrodes in the bath near the surface of the oocyte and zero their offset potentials. Impale the oocyte with the voltage electrode and then the current electrode (see Note 6). Clamp the oocyte at the resting membrane potential (40 mV) and ensure the leak current is as close to zero as possible. 6. Record photocurrents using the stimulus protocol described previously [13]. In brief, clamp the membrane at potentials ranging from 100 mV to +40 mV in +20 mV steps, with a 1.5 s illumination period at each step. Record photocurrents in sodium and potassium bath solutions first and then in calcium bath solution. The same oocyte can be used to measure photocurrents in all four bath solutions before switching to another oocyte. When recording in calcium bath solution, visually inspect the photocurrent traces to ensure the intracellular calcium is effectively buffered by BAPTA and that the endogenous chloride channels are not being activated as this would impact the validity of the data. Figure 3 illustrates the typical appearance of channelrhodopsin photocurrent traces in the case of fully suppressed, unsuppressed, and partially suppressed calcium-activated chloride currents. If chloride currents are present as in Fig. 3b or c, add more BAPTA-AM to the oocyte media and/or increase incubation time until chloride currents are abolished. 7. When finished recording with one oocyte, switch off voltage clamp mode and raise the electrodes out of the bath solution. Discard the oocyte and flush the chamber with sodium solution to prepare for the next oocyte. 8. Repeat steps 4–7 with a new oocyte. 3.4 Data Processing, Permeability Ratio Calculations, and Kinetic Analysis

The gating kinetics of channelrhodopsin-2 and the C1C2 chimera are best described using a four-state model with two open states and two closed states [14–17]. A schematic representation of this model is given on the left-hand side of Fig. 4. The cycle begins in the first closed state, C1. In this state, ChR is fully dark-adapted and no ionic current is flowing through the channel pore. Upon irradiation with blue light indicated by the blue lightning bolt, the covalently bound all-trans retinal located in the center of the protein undergoes a trans to cis isomerization reaction. This triggers a series of conformational changes in the protein that lead to the first open state, O1, which is a high-conducting open state where maximal current flows through the open pore. The transition rate from C1 to O1 is defined by the rate constant k1. Upon extended illumination with blue light, the population of activated channels will

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Fig. 3 Representative photocurrent traces of channelrhodopsin expressed in X. laevis oocytes in Ca2+ bath solution. Blue bars indicate channel activation via irradiation with 470 nm LED light. The voltage step protocol is depicted in the subplot. (a) Isolated calcium photocurrent through C1C2 in an oocyte treated with BAPTA-AM for 1 h where the intracellular calcium is buffered effectively and endogenous chloride current is sufficiently suppressed. (b) Current recorded in calcium solution from an oocyte expressing C1C2 that was not treated with BAPTA-AM. Here, intracellular calcium is not buffered, and a large contaminating outward chloride current can be seen upon photoactivation when the membrane is clamped at Vm ¼ 100 mV and 80 mV. (c) Current recorded through C1C2 in an oocyte treated with BAPTA-AM for 30 min where the intracellular calcium is only partially buffered due to inadequate BAPTA loading time. The activation of the contaminating outward chloride current visible for the Vm ¼ 100 mV trace is delayed compared to the recording in (b), which indicates partially suppressed chloride channel activation

shift between the first high-conducting open state (O1) and a second, lower-conducting open state (O2) until an equilibrium is reached. The transition rates associated with the transitions from O1 to O2 and from O2 to O1 are given by the rate constants e12 and e21, respectively. When the light is turned off, the channels close by transitioning to either the first closed state, C1, or a second closed state, C2. The rates for the transitions from O1 to C1 and from O2 to C2 are given by the constants Gd1 and Gd2, respectively. The second closed state is a desensitized, or light-adapted state. Channels activated from C2 will undergo a transition to the second, lower-conducting open state, O2, and is described by the rate constant k2. Lastly, channels in the C2 state will spontaneously transition back to the first closed state (C1) in the dark to complete the cycle. The rate of the transition C2 to C1, also called the recovery rate, is given by Gr. The rates of each of these transitions can be estimated from the photocurrent data collected from TEVC experiments. Since TEVC gives whole-cell recordings, the resulting photocurrent trace is a measure of the sum of the flow of ions through each individual ChR channel populating the cell membrane in either of the two open states. Therefore, some key features of the photocurrent curve can be defined and related to specific steps in the photocycle (Fig. 4, right). The peak current (Ip) is the maximum current amplitude

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Fig. 4 Kinetics of ChRs. (a) Schematic representation of the four-state photocycle of channelrhodopsin. Blue lightning bolts indicate light-dependent processes. Arrows indicate transitions between states and are labelled with their corresponding rate constants. (b) Representative photocurrent trace of channelrhodopsin measured via TEVC. The blue bar represents channel activation by irradiation with 470 nm light. Kinetic parameters and key features of the curve used for kinetic and ion selectivity analysis are defined

reached upon channel activation. The portion of the curve beginning from the time the light is turned on to peak current can be attributed to the sum of the transitions from the two closed states to the two open states. The time constant, τ on, is therefore proportional to the sum of the rate constants k1 and k2. Under continuous illumination, the current then decays from peak current to a constant amplitude which is referred to at the steady state, or stationary, current (Iss). This portion of the curve from Ip to Iss is related to the sum of the forward and reverse transitions between the two open states. Therefore, the time constant τ decay is proportional to the sum of the transition rates e12 and e21. Once the light is turned off, the channels close, transitioning from O1 to C1 and from O2 to C2, and the measured current undergoes a rapid exponential decay to zero. Similar to the other two time constants, the off current represents the sum of these two transitions, and its time constant, τ off, is proportional to the sum of Gd1 and Gd2. The rate constants of each individual transition can be estimated via computational modeling of the kinetic parameters in a program

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such as MATLAB. However, we describe a simplified analysis in this work and refer the reader to Prignano et al., for a more in-depth and computation-intensive method [17]: 1. All data processing and analysis can be performed in either pCLAMP software or MATLAB. First, subtract any leak current from all traces. The leak current is the passive diffusion of ions through minor imperfections in the cell membrane. A healthy oocyte will have a leak current 1000 proteins in coupled networks [11], and the challenges faced by the pharmaceutical industry in finding new medicines that target synaptically mediated diseases, there is a strong need for techniques to measure synaptic function at a scale that will enable drug discovery. Despite this need, existing methods for measuring synaptic signaling are inadequate, often due to limitations of either throughput or information content. Manual patch clamp measurements are

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Gabriel B. Borja, Himali Shroff, and Hansini Upadhyay contributed equally to this chapter.

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the gold standard in electrophysiology but have limited throughput, and automated patch clamp is presently incompatible with synaptically connected neurons. Multi-electrode array (MEA) recordings measure network activity, but require custom, expensive plates, which can limit their use in high-throughput applications, and cannot assign recordings to individual cells study effects on neuronal subtypes. For these reasons, there remains a need for robust, scalable assays of synaptic function. Optogenetics has emerged over the last decade as a powerful research tool that can potentially bridge the gap between highthroughput assays of synaptic function and drug discovery for neurological disorders. The hallmark of optogenetics is precise optical control of neuronal activity through genetically targeted, light-gated ion channels expressed in neurons [12]. These channels can be co-expressed in neurons along with reporters of key physiological readouts, such as voltage or calcium, to enable all-optical methodologies for measuring neuronal excitability, synaptic transmission, and network activity. Here we describe two optogenetic assays that can be used to measure synaptic signaling with high fidelity. The first assay is a pre-synaptic calcium assay that utilizes the channelrhodopsin, CheRiff [13], and the calcium reporter jRGECO1a [14] fused to the C-terminus of synaptophysin. Through targeted optogenetic stimulus, pre-synaptic changes in calcium influx can be measured. The second assay enables measurement of synaptic transmission. To achieve this readout, CheRiff and cytosolic jRGECO1a are expressed in non-overlapping sets of neurons using a Cre recombinase approach that is described below. Pre-synaptic neurons express CheRiff, and post-synaptic neurons express the calcium indicator jRGECO1a. Pharmacological validation data is provided in each case. The chapter provides detailed descriptions of the required methodologies to perform the two assays as well as additional notes to highlight potential pitfalls and solutions.

2

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2.1 Concept and Optogenetic Constructs

To probe synaptic function with an all-optical assay that can be scaled to high throughput, we use two genetically encoded components. The blue light-gated cation channel CheRiff [13] is used to trigger neuronal action potentials, and a yellow light excited calcium sensor jRGECO1a [14] is used to track changes in calcium levels. CheRiff is a channelrhodopsin that produces relatively large currents in response to low blue light levels, and jRGECO1a is a fast and sensitive protein-based sensor that can reliably detect the calcium influx induced by a single neuronal action potential. When pairing these optogenetic tools, reducing optical crosstalk is critical

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5 mm, that excites jRGECO1a fluorescence (red). 380 nm UV light and blue stimulation light from LEDs are reflected off a DMD, which is imaged onto the sample, enabling patterned illumination that can be rapidly reconfigured as a function of space and time. In the “Oreo” geometry, blue stimulus is applied on either side of the recording region. (b) Rat hippocampal neurons lentivirally transduced with constructs CheRiffEBFP2 (blue) and jRGECO1a (red), both expressed under the hSYN promoter. Here the UV light, which illustrates the stimulus pattern, excites fluorescence from CheRiff-EBFP2. The stimulus light, which does not overlap with the 570 nm excitation light, illuminates ~1.7-mm-high strips above and below the recording region

to success. The 570 nm yellow light used to excite jRGECO1a only minimally excites CheRiff, so optical recordings have only minor effects on neuronal behavior [15]. In contrast, jRGECO1a markedly changes its fluorescent brightness in response to the blue light used for stimulation [14, 15]. To avoid optical crosstalk issues, we pattern the illumination light on the Firefly microscope [16]. The 570 nm laser used to excite jRGECO1a passes through horizontal and vertical slits that are imaged onto the sample, providing a tightly bounded rectangular illumination area (Fig. 1a). The 470 nm light-emitting diode (LED) light for stimulating CheRiff is patterned to illuminate rectangular regions above and below the 570 nm rectangle using a digital micromirror device (DMD) (Fig. 1a). In this way, the blue and yellow illumination regions are not overlapped (Fig. 1b), minimizing crosstalk issues. In the future, we hope that new calcium sensors operating in the red spectrum

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Fig. 2 Pre-synaptic calcium assay constructs and images. (a) The transmembrane protein CheRiff-EBFP2 and pre-synaptically targeted Syp-jRGECO1a are expressed in all transduced neurons under the hSYN promoter. TS trafficking sequence. EBFP2 enhanced blue fluorescent protein 2. Pre-synaptic trafficking of the calcium sensor is achieved by fusing the sensor onto the C-terminus of synaptophysin, targeting jRGECO1a to the outer leaflet of synaptic vessicles. (b–d) Primary E18 rat hippocampal neurons co-cultured with rat glial cells in Ibidi 96-well plates. Images acquired 12 days after plating and 6 days after transduction. (b) Synaptophysin-jRGECO1a, a pre-synaptically targeted calcium sensor under control of the hSYN promoter demonstrating sensor localization in synapses (small puncta). (c) CheRiff-TS-EBFP2, a blue light-gated ion channel under control of hSYN promoter, showing membrane localization. (d) Merge of (b) (red) and (c) (blue). Fluorescence images captured with a 40
oil objective, NA ¼ 1.3

such as K-GECO1 [17] or NIR-GECO1 [18] may have minimal optical crosstalk, obviating the need for light patterning and thus simplifying experiments. We have cloned the DNA constructs encoding both the calcium sensor and the optogenetic actuator into the FUGW lentiviral backbone plasmid [19] and use the second generation lentiviral packaging plasmids [20] for viral particle production. With lentiviral transduction and expression driven by the human SYNAPSIN I (hSYN) promoter [21], we can selectively express these constructs at high levels in the majority of neurons with minimal expression in the co-cultured glial cells. For each of the constructs (e.g., Fig. 2a), the illustrated cassette is inserted between the hSYN promoter and the WPRE posttranscriptional regulatory element [22], which enhances protein expression levels.

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The pre-synaptic calcium assay (Fig. 2a) uses two constructs. The channelrhodopsin CheRiff is fused with the trafficking sequence from KIR 2.1, which improves mammalian membrane trafficking [23], and the blue fluorescent protein EBFP2 [24]. EBFP2 enables visualization of protein trafficking (Fig. 2c) and leaves the EGFP channel available for other fluorescent sensors or cell labeling. The jRGECO1a calcium reporter is trafficked to the pre-synaptic compartment by fusing it to the C-terminus of synaptophysin (Syp-jRGECO1a), a protein expressed in the membrane of pre-synaptic vesicles. Syp-jRGECO1a expression is enriched in puncta that localize along CheRiff-expressing dendrites (Fig. 2b, d), suggestive of synaptic localization. Using optical sectioning with a confocal microscope, Syp-jRGECO1a also co-localizes with pre-synaptic terminals, as labeled by an α-synaptophysin antibody, that appear to be synapsed onto the cell body and proximal dendrites, as labeled by MAP 2 (Fig. 3). Syp-jRGECO1a trafficking is not perfectly localized to the pre-synaptic terminals, as there can be some expression in compartments in the cell body and along neurites, so care must be taken to account for these contributions to calcium signaling. If recordings are performed at high magnification, regions too large to correspond to pre-synaptic boutons can be excluded by morphological size filtering. Alternatively, L-type calcium channels Cav1.2 and Cav1.3, which dominate cytosolic B

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Fig. 3 Synaptic localization of pre-synaptic calcium sensor. Hippocampal neurons from E18 rats cultured for 14 days, fixed, and immunolabeled. Images were collected with a 60
objective on a confocal microscope. (a) Antibody-labeled microtubule-associated protein 2 (MAP 2), which stains the cell body and proximal dendrites. (b) Antibody-labeled synaptophysin, a protein localized to the membrane of pre-synaptic vesicles. (c) Calcium sensing protein jRGECO1a fused to synaptophysin. (d) Overlay of jRGECO1a and synaptophysin shows many overlapping puncta. (e) Overlay of synaptophysin and MAP 2 shows pre-synaptic puncta along the dendrites and cell body. (f) Overlay of jRGECO1a and MAP 2 shows puncta along the dendrites and cell body in the same locations as synaptophysin, demonstrating the pre-synaptic localization of the sensor

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Fig. 4 Pre-synaptic calcium assay functional data. (a) The fluorescent Syp-jRGECO1a signal calculated by taking the flat average of fluorescence across the full record regions. Calcium transients were detected in response to optogenetic stimuli (5 ms, 125 mW/cm2 blue light pulses). We have previously measured these stimuli to trigger action potentials with high fidelity; 125 mW/cm2 will open the majority of CheRiff channels [13], strongly depolarizing neurons. Action potentials from the stimulated region will propagate into the recording region and induce calcium influx to the pre-synaptic terminals. Calcium enters the cell through voltage-gated calcium channels, predominantly Cav2.1 and 2.2, which are selectively blocked by toxins ATX and CTX, respectively. ATX: 0.5 μM ω-agatoxin IVA. CTX: 1 μM ω-conotoxin GVIA. (b) Recordings were made using both the pre-synaptically targeted jRGECO1a and a cytosolic (untargeted) jRGECO1a. Recordings as shown in (a) were made before (pre) and after (post) toxin addition and integrated as a measure of total calcium influx. Post-signals were normalized to pre-signals to minimize effects of well-to-well variability, and that ratio was normalized to the vehicle condition

calcium transients during an action potential in central nervous system neurons, can be pharmacologically blocked with isradipine and nimodipine [25, 26]. Figure 4 shows calcium transients recorded with the pre-synaptic calcium assay. Optogenetic stimuli induce action potentials, which trigger calcium influx through voltage-gated calcium channels. Because of the patterned optogenetic illumination (Fig. 1), electrical signals must propagate down neurites from the stimulus region into the recording region for calcium influx to be detected. This minimizes optical crosstalk (direct blue light modulation of jRGECO1a brightness) and any undesired signal from calcium influx directly through the nonselective cation channel CheRiff. In pre-synaptic boutons, calcium enters the cell predominantly through P-type and N-type voltage-gated calcium channels Cav2.1 and Cav2.2 [27–29], which are transiently opened by action potentials. Figure 4a shows the marked reduction in calcium transient amplitude by block of Cav2.1 by ATX (ω-agatoxin IVA), Cav2.2 block by CTX (ω-conotoxin GVIA), and co-block with both toxins. Figure 4b compares block in pre-synaptic boutons and in the cytosol. Cav2.1 and 2.2 contribute to calcium influx in both compartments but account for a greater portion of the calcium signal in the boutons. The expected pharmacological responses and accurate sensor trafficking to pre-synaptic terminals build confidence in the

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assay for characterizing the effects of Cav-modulating compounds under development or for phenotypic drug screening for synaptic modulators. 2.3 Synaptic Transmission Assay

The synaptic transmission assay is implemented by expressing the optogenetic actuator CheRiff in a subset of cells and the calcium sensor, cytosolic jRGECO1a, in a non-overlapping set of cells (Fig. 5). Blue light stimulation triggers action potentials in pre-synaptic, CheRiff-expressing neurons, and synaptically mediated calcium signals are detected in post-synaptic, jRGECO1a-expressing neurons. Expression in non-overlapping sets of neurons is implemented with the Cre recombinase/lox system [30], which has been used previously for assaying synaptic signaling [31]. In the “CreOFF” vector, CheRiff is placed between two lox sites oriented in the same direction [30] (Fig. 5a). CheRiff is expressed in the absence of Cre recombinase, but when Cre is present, it excises the open reading frame (ORF) and shuts down expression (Fig. 4b). In the “CreON” vector, jRGECO1a is placed in reverse orientation between two pairs of opposed lox sites, the flip-excision (FLEX) switch system [30]. The backward construct is not expressed in the absence of Cre, but when Cre is expressed, the jRGECO1a is flipped, and the extraneous lox sites are excised, permanently activating expression (Fig. 5b). To obtain a balance of CheRiff- and jRGECOa1-expressing neurons, lentiviral particles expressing Cre recombinase are added at low multiplicity of infection, so that stochastically, approximately half of the neurons in the culture express Cre and half do not. Neurons that do not express Cre express CreOFF-CheRiffEBFP2 only, while neurons that express Cre express CreONjRGECO1a but not CheRiff. Figure 5c–e shows expression of CheRiff and jRGECO1a in nearly non-overlapping sets of neurons. The platform is efficient, with >90% of neurons expressing exactly one of the two constructs (Fig. 5e). Figure 6 shows functional recordings in the synaptic transmission assay. A series of optogenetic stimuli trigger action potentials in the pre-synaptic neurons, which induces calcium transients in the post-synaptic neurons via synaptically mediated signaling (Fig. 6a). By titrating the dose of Cre recombinase, the ratio between preand post-synaptic neurons can be smoothly tuned across the full range (Fig. 6c). When the number of pre- and post-synaptic neurons is highly imbalanced, the amplitude of the post-synaptic calcium transients is reduced (Fig. 6d). The synaptic transmission assay responds as expected to tool compounds (Fig. 6a, b). Block of the receptors for excitatory and inhibitory neurotransmitters almost eliminates the post-synaptic calcium transients (purple). Similarly, block of Cav2.1, 2.2, and 2.3, which strongly attenuates pre-synaptic calcium (Fig. 4) and thus pre-synaptic vesicle release, almost completely blocks the

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Fig. 5 Synaptic transmission assay concept and images. (a) Three constructs for probing synaptic transmission: Cre recombinase, CreOFF-CheRiff, and CreON-jRGECO1a. Green arrowhead: loxFAS. Black arrowhead: lox2272. White arrowhead: loxP. (b) Low-dose Cre stochastically transduces a subset of neurons, leading to CheRiff and jRGECO1a expression in non-overlapping sets of cells. In pre-synaptic, Cre() neurons, CheRiff is expressed, but the reversed jRGECO1a is not. In post-synaptic, Cre(+) neurons, a circular piece of DNA containing the CheRiff expression is excised and silenced, while the jRGECO1a cassette is flipped and activated. (c–e) Fluorescence images through a 10x air objective (NA ¼ 0.45) of primary hippocampal neurons from E18 rats cultured for 12 days with a 30 nL dose of Cre-lentivirus. (c) CreON-jRGECO1a, a somatic calcium sensor under control of the hSYN promoter. Image corrected (γ ¼ 0.2) to highlight dim neurites. (d) CreOFF-CheRiff-EBFP2 under control of the hSYN promoter. Image corrected (γ ¼ 0.3) to highlight dim neurites. (e) Merge of (d) (red) and (e) (blue) showing that the constructs are predominately expressed in distinct populations

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Fig. 6 Synaptic transmission assay functional data. Recordings are made in primary E18 rat hippocampal neurons cultured for 14 days. (a) Post-synaptic calcium transients, calculated by taking the flat average over the full record region, in response to a patterned optogenetic stimulus pulses of 5 ms at 125 mW/cm2. The calcium signal is modulated by tool compounds. 1 μM PMA (phorbol-12-myristate-13-acetate) is an enhancer of pre-synaptic vesicle release. “Post-synaptic block” is a mixture of three neurotransmitter receptor blockers: 20 μM gabazine blocks the GABAA receptor, 100 μM NBQX blocks the NMDA receptor, and 50 μM AP-V blocks the AMPA receptor. “Pre-synaptic CaV block” is a mixture of three toxins that block the voltage-gated calcium channels that are enriched at pre-synaptic boutons: 0.5 μM ω-agatoxin IVA blocks Cav2.1, 1 μM ω-conotoxin GVIA block Cav2.2, and 1.2 μM SNX-482 blocks Cav2.3. (b) Recordings as shown in (a) were made before (pre) and after (post) compound addition and integrated over time as a measure of total post-synaptic activity. Post-signals were normalized to pre-signals to minimize effects of well-to-well variability, and that ratio was normalized to the vehicle condition. Activation and strong block were observed. (c) The fraction of Cre(+) and Cre() neurons as a function of Cre lentiviral dose was assessed using CreON-nucTagRFP (nuclear-localized red fluorescent protein) and CreOFF-nucEGFP (nuclear-localized green fluorescent protein). Morphological image segmentation was used to identify each fluorescent nucleus and identify it as “red,” “green,” or both. By tuning the Cre dose, we can accurately control the number of CreON- and CreOFF-expressing neurons, always with minimal co-expression. (d) The integrated calcium signal (black trace in (a)) as a function of Cre dose. As expected, the total amount of synaptic signaling is largest when there is a somewhat balanced mixture of CheRiff- and jRGECO1a-expressing neurons

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signal. In contrast, the phorbol ester phorbol-12-myristate-13-acetate (PMA), which has been shown to enhance pre-synaptic vesicle release [32, 33], increases post-synaptic calcium transient amplitude. Thus, the assay can detect a variety of expected modulators and can be deployed in drug discovery or basic science.

3

Materials

3.1 Lentivirus Production Materials

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HEK293T cells (ATCC #CRL3216).

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Lentiviral Production Medium 1 (see recipe).

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Lentiviral Production Medium 2 (see recipe).

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PBS (phosphate-buffered saline) (1.06 mM KH2PO4, 155.17 mM NaCl, and 2.97 mM Na2HPO4-7H2O)

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Freezing medium: 20% dimethyl sulfoxide (DMSO) in fetal bovine serum (see recipe).

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Second- or third-generation lentiviral plasmid encoding optogenetic construct.

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Second-generation viral packaging mix containing plasmids for PsPAX2 and PMD2.G (contains VSVG gene), supplied as 250 μg in a 0.5 μg/mL solution (Cellecta, Cat# CPCP-K2A).

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Opti-MEM reduced serum medium (ThermoFisher Scientific, Cat# 31985-070).

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Polyethylenimine (PEI) MAX40000 transfection reagent (see recipe).

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Lenti-X Concentrator (Takara Clontech, Cat# 631231).

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Neurobasal medium (ThermoFisher Scientific, Cat# 10888022).

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Lenti-X GoStix (Takara Clontech, Cat# 631243).

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Lenti-X qRT-PCR Titration Kit (Takara Clontech, Cat# 631235).

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10 cm (diameter) tissue culture dishes.

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15 cm (diameter) tissue culture dishes.

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15 mL conical tubes.

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50 mL conical tubes.

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1.8 mL cryovials.

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500 mL 0.45 μm filter units.

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0.6 mL sterile tubes.

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10-layer HYPERFlask (Corning Cat# 10030).

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500 mL conical centrifuge tube.

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750 mL polypropylene bottle.

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3.1.1 Lentiviral Production Medium 1

To prepare 500 mL of medium, combine 440 mL of high-glucose DMEM, 5 mL of GlutaMAX (Life Technologies, Cat# 35050061), 5 mL of nonessential amino acids, and 50 mL of heatinactivated fetal bovine serum (HyClone, Cat# SH30071.02HI). Mix and vacuum filter inside the biosafety cabinet. The addition of penicillin/streptomycin is not recommended as it may reduce virus yield. Recommended shelf life is 14 days at 4  C.

3.1.2 Lentiviral Production Medium 2

Add 5 mL of penicillin/streptomycin to Medium 1. Recommended shelf life is 14 days at 4  C.

3.1.3 PEI Transfection Reagent

We use a linear chain polyethylenimine (PEI) with a characteristic molecular weight for the free base of 22 kDa (Polysciences, cat.no. 24765-2). A stock solution is prepared at 1 mg/mL using molecular biology grade water by placing 10 mg of PEI in 9 mL water. Bring the PEI solution to pH of 7.10 using 1 M sodium hydroxide (NaOH), as the solution is initially acidic (pH 2.0–3.0). Bring the final volume to 10 mL to obtain a concentration of 1 mg/mL. Filter the solution using 0.22 μm filter, aliquot, and store at 80  C.

3.1.4 Freezing Medium

Add 8 mL DMSO (Sigma-Aldrich, Cat# D2650-100ML) to 32 mL FBS (HyClone, Cat# SH30071.02HI) to achieve a final DMSO concentration of 20%. Sterile filter before use.

3.2 Neuronal Culture Materials

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E18 Rat Tissue (BrainBits, Cat# SDEhp).

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Poly-D-lysine hydrobromide (Sigma-Aldrich, Cat# P640710X5MG).

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96-well plates (μ-Plate 96 Well, #1.5 polymer coverslip, ibiTreat tissue culture-treated, sterile).

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Papain enzyme with Hibernate E-CA media (BrainBits, Cat# PAP/HE).

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P1 Rat Cortex (BrainBits, Cat# PRcx Rat).

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Rat neuronal culture medium (see recipe).

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Rat glial culture medium (see recipe).

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1
PBS (phosphate-buffered saline) (1.06 mM KH2PO4, 155.17 mM NaCl, and 2.97 mM Na2HPO4-7H2O).

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0.25% trypsin-EDTA (Sigma-Aldrich, Cat# T-4049-100ML).

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Poly-D-lysine-coated T-175 flasks (ThermoFisher Scientific, Cat# 132705).

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Dimethyl sulfoxide (DMSO; Sigma-Aldrich, Cat# D2650100ML).

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1.8 mL cryovials.

l

Heat-inactivated FBS fetal bovine serum (HyClone, Cat# SH30071.02HI).

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3.2.1 Rat Neuronal Culture Medium

To make 500 mL of media, mix 475 mL Neurobasal-A medium (ThermoFisher Scientific, Cat# 10888022), 5 mL GlutaMAX (ThermoFisher Scientific, Cat# 35050061), 5 mL penicillin streptomycin, 10 mL 50
B-27 Supplement (ThermoFisher Scientific, Cat# 17504-044), and 5 mL 100
N2 Supplement (ThermoFisher Scientific, Cat# 17502048). Recommended shelf life is 14 days at 4  C. Immediately prior to use with primary rat neuronal cultures, add mouse recombinant laminin at a final concentration of 1 μg/ mL (ThermoFisher Scientific, Cat# 23017-015) and the neurotrophic factors recombinant BDNF (R&D Systems, Cat# 248-BDB) and recombinant GDNF (R&D Systems, Cat# 212-GD); each neurotrophic factor should be added to a final concentration of 10 ng/mL.

3.2.2 Rat Glia Culture Medium

To make 500 mL of medium, add 5 mL penicillin streptomycin to 500 mL NbASTRO medium (BrainBits, Cat# NbASTRO). Recommended shelf life is 14 days at 4  C.

3.3 Optical Imaging Materials

Tyrode’s has low autofluorescence and a stable pH under ambient atmosphere. To prepare, make a solution in molecular biology grade water with the following composition: 10 mM HEPES, 125 mM NaCl, 2 mM KCl, 3 mM CaCl2, 1 mM MgCl2, and 30 mM glucose. Set the buffer pH to 7.35  0.05.

3.3.1 Tyrode’s Imaging Buffer

4

Methods

4.1 Production of Lentiviral Particles Encoding Channelrhodopsin (CheRiff) and Protein-Based Fluorescent Calcium Sensor (jRGECO1a)

Lentiviral particles can drive high and stable expression levels as well as transduce post-mitotic cells such as cultured neurons with high efficiency [20, 22, 34–36] and minimal cell toxicity. In our experience, lentiviral transduction efficiency surpasses ~90% of cultured neurons. We previously published on lentivirus production in 15 cm dishes [37]. In order to obtain higher quantities and reduce batch-to-batch variability, we have optimized our lentiviral production methods in 10-layer HYPERFlasks. Compared to 15 cm dishes, HYPERFlasks have an eight-fold larger area and a ten-fold larger volume and produce 11-fold more viral genome copies and 30-fold more functional virus particles. Detailed below are the steps for lentivirus production: culture of HEK293T (293T) cells, transfection of 293T cells with viral packaging plasmids, virus harvest, virus concentration, and virus quality control (QC). Overall Timeline: Day 0—Thaw first round of 293T cells onto 2
10 cm dishes. Day 3—Passage cells and make 2
15 cm dishes. Day 6 or 7—Passage cells 1:6 and make 12
15 cm dishes or passage one 15 cm dish into one HYPERFlask (this is a 1:8 split) (see Note 1).

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Day 10—Transfect cells and express for 48–60 h. Day 12+—Harvest the virus. Unless otherwise stated, all virus production and cell culture work must be conducted under sterile conditions inside a biological safety cabinet using BSL2 safety requirements and with cultures maintained in biological incubators at 37  C in 5% CO2. 4.1.1 Preparing Cryo-Stocks of 293T Cells for Virus Production

1. To minimize batch-to-batch variability in the production of different lentiviral stocks, a large cryobank of low-passage 293T stocks can be created which can be used to initiate multiple rounds of virus production. To prepare these cryostocks, thaw a cryovial of 293T cells in a 37  C water bath for 3 min in preparation for culture in a 10 cm tissue culture dish. 2. Transfer the contents of the vial to a 15 mL conical tube, and rinse the cryovial with 1 mL of Lentiviral Production Medium 2 (see recipe in Materials section above) to collect any remaining cells. Transfer this suspension dropwise to the 15 mL conical tube, and gently swirl the tube to avoid osmotic shock to the thawing cells. 3. Add an additional 8 mL of Lentiviral Production Medium 2, bringing the total volume to 10 mL. 4. Centrifuge the cell suspension for 5 min at 300
g, room temperature (RT), to pellet the cells. 5. Aspirate the supernatant to remove all DMSO-containing freezing solution; then resuspend the cell pellet in 1 mL of Lentiviral Production Medium 2 by gently pipetting up and down at least 5–10 times with a P-1000 μL pipettor. 6. Bring the volume to 10 mL with Lentiviral Production Medium 2, and plate onto a 10 cm tissue culture dish. 7. Passage 293T cells every 3 days with Lentiviral Production Medium 2 when cells reach 80–90% confluency using the method described in Subheading 3.1.2 (below). 8. From the 1
10 cm tissue culture dish, 293T cells are passaged onto 2
15 cm tissue culture dishes. Once in a 15 cm tissue culture dish, the 293T cells can be passaged every other day with a 1:4 dilution factor (i.e., the cells from 1
15 cm dish can be dissociated and plated onto 4
15 cm dishes for expansion). 9. Once 32
15 cm dishes are prepared, the cells are dissociated, counted, and frozen in cryovials. On average, approximately five million 293T cells are obtained per 15 cm tissue culture dish. These cells can be evenly divided and frozen in two cryovials. This step involves preparing a cell suspension of ten million cells per mL in Lentiviral Production Medium 2 and

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adding 500 μL of this suspension to a cryovial, followed by dropwise addition of 500 μL of freezing medium (see recipe in Materials section above), bringing the total DMSO concentration to 10%. The cryovials are transferred immediately into a 80  C freezer in a ~1  C/min isopropyl alcohol freezing container. After 16–24 h, cryovials are transferred to a liquid nitrogen storage unit. 4.1.2 Passage 293T Cells for Virus Production

1. From a dish with ~80% confluent 293T cells, aspirate the medium from the dish, and then gently rinse with a sufficient volume of PBS to cover the entire surface of the dish (6 mL for a 10 cm dish and 20 mL for a 15 cm dish). Aspirate the PBS. 2. Dislodge the cells from the surface by the vigorous addition of 8 mL of PBS with a 10 mL serological pipette, rinsing several times until the cells visibly lift from the surface. No peptidase is required. 3. Transfer the cell suspension to a 50 mL conical tube. Wash the dish surface to collect residual cells, and add to the 50 mL conical tube. 4. Pellet the cells by centrifuging 5 min at 300
g, RT. 5. Aspirate the supernatant and resuspend the cells in 1 mL of Lentiviral Production Medium 2. 6. Pool the cells from different plates to homogenize density in the next round. 7. Add 1 mL of the pooled cell suspension to 500 mL of Lentiviral Production Medium 2, and pour the contents to a 10-layer HYPERFlask. Add additional media to the HYPERFlask up to the top thread of the screw cap. Transfer the HYPERFlask to the incubator. If splitting cells onto 15 cm dishes, add the 1 mL cell suspension to 24 mL of Lentiviral Production Medium 2 and plate (see Note 2).

4.1.3 Transfect 293T Cells

Transfect 293T cells in 10-layer HYPERFlask when they reach a confluency of 80–90% (Fig. 7a). 1. To a 50 mL conical tube, add 160 μg of plasmid containing the gene of interest (see Note 3), 200 μg of the mixture of packaging and envelope plasmids PSPax2 and pMD2.G, 4 mL of Opti-MEM serum-free medium, and 1.44 mL of 1 mg/mL PEI transfection reagent (see recipe in Materials section above). 2. Mix thoroughly and incubate for 10 min at RT. 3. Add 500 mL of Lentiviral Production Medium 1 (see recipe in Materials section above; see Note 4) to DNA/PEI mixture. Mix well by inverting several times gently. Pour out existing

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media from the HYPERFlask, and slowly replace with the 500 mL transfection mixture. Top off the HYPERFlask with Medium 1. 4. Culture the transfected cells in cell incubator for an additional 48–60 h before harvesting the lentiviral particles. 4.1.4 Harvest Lentiviral Particles

Lentivirus is a Biosafety Level 2 (BSL2) agent that can infect human cells. It is recommended that, in addition to the standard gloves and lab coats, sleeves be worn and that all containers and pipettes in contact with the virus be bleached prior to disposal in the biohazard waste containers. For 293T cells transfected with a fluorescent protein-expressing construct, transfection efficiency can be confirmed using a fluorescence microscope (Fig. 7b–d). After 24 h, 30–50% of 293T cells should exhibit fluorescence, indicating good PEI transfection efficiency. In 293T cells, a universal CMV promoter drives expression of the full RNA transcript that gets packaged into the viral capsids, so protein expression can be detected even if the promoter proximal to the optogenetic construct is not expressed in 293T cells. If the proximal promoter is expressed in HEK cells, after 48–60 h, expression efficiency should exceed 80% as released virus transduces additional cells. Poor transfection efficiency is indicative of a production batch with lower viral titer. Figure 7b–d shows transfection efficiency for a typical batch. 1. Pour the lentivirus-containing supernatant from each HYPERFlask into a 500 mL conical tube. 2. Centrifuge 5 min at 2000
g at RT, to precipitate any HEK cells or debris. 3. Filter the supernatant through a 500 mL, 0.45 μm filter unit to remove residual cell debris.

4.1.5 Concentrate and Characterize Lentivirus Stocks

To minimize the volume of lentiviral reagent delivered to neuronal cultures, reduce required storage space, and remove any potentially deleterious components in the 293T cell culture supernatant, the virus is concentrated 30-fold and resuspended in neuronal culture medium. 1. Mix the filtered virus suspension with the Clontech Lenti-X Concentrator reagent, and centrifuge in 750 mL polypropylene bottle according the manufacturer’s protocol. 2. Resuspend the virus-containing pellet from the Lenti-X concentration in Neurobasal medium to increase viral concentration 30-fold relative to the supernatant, e.g., the viruscontaining pellet obtained from 540 mL of supernatant is resuspended in 18 mL of Neurobasal medium (see Note 5).

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Fig. 7 Virus production and characterization. (a) A phase contrast image of HEK-293T cells immediately before PEI transfection. The confluency shown is on the high end of the optimal range. (b) A phase contrast image of 293 cells 60 h after PEI transfection with the lentiviral plasmids. The expression cassette encodes TagRFP expressed under the human synapsin promoter. (c) A fluorescence image of TagRFP from the same region. (d) An overlay, showing high transfection efficiency. (e, f) Quantification of viral genome concentration through quantitative polymerase chain reaction (qPCR). (e) Using a standard reference with known concentration, we perform a log-linear fit on the dilution series to calibrate the relationship between cycle number and concentration of viral genomes. (f) We perform a dilution series on the new virus batch to be titered and perform qPCR, converting cycle number to concentration using the fit in (e). The intercept of the fit line is the estimated virus stock concentration

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3. Distribute the concentrated stocks into sterile 0.6 mL microcentrifuge tubes, and store at 80  C (see Note 6). 4. The lentivirus stocks can be qualitatively titered with Clontech Lenti-X GoStix and quantitatively titered with Clontech LentiX qRT-PCR using the manufacturer’s protocols. Viral genome titers of 109 to 1010 copies/mL are normally achieved (Fig. 7e, f) in the 30-fold concentrated stocks. The functional titer, quantified by single-copy transduction efficiency at high dilution factors, is around 50 times lower at ~108 IFU/mL. With a typical dose of 0.3 μL/well of a 96-well plate, the 18 mL of 30
stock can transduce 60,000 wells. 4.2 Preparation of Primary Rat Hippocampal Neurons for Optogenetic Measurements

Cultures of primary rat hippocampal neurons display robust action potential firing and synaptic activity. They express the engineered constructs at high levels, providing high sensitivity to optogenetic stimulation with blue light and high signal-to-noise ratio (SNR) calcium recordings with yellow light excitation [13, 15, 31, 37]. Co-culture with glial cells improves neuronal health and enhances synaptic connectivity. Below we describe protocols for the culture and cryobanking of rat glial cells, the co-culture of neurons and glia, and lentiviral transduction of cultured neurons with optogenetic constructs. Overall Timeline: 1. Day in vitro 0 (DIV): Plate E18 rat hippocampal neurons dissociated from tissue along with rat glial cells. 2. DIV 6: Deliver lentiviral constructs encoding optogenetic protein and calcium sensor. 3. DIV 13–16: Optogenetic stimulation and calcium imaging.

4.2.1 Preparation, Culture, and Cryobanking of Primary Rat Glial Cells

The rodent glial protocols are adapted from De Giorgio et al. [38] and BrainBits, LLC dissociation protocols (http://www. brainbitsllc.com/postnatal-cortex-plating-protocol/). The cortex from P1 post-natal rat pups is ordered from BrainBits; cells from three cortices (left and right hemispheres) are plated onto one polyD-lysine-coated T-175 flask. The cortices are dissociated following section 3.2.3. Dissociation of Rat Hippocampal Tissue with two changes: (1) DNase is added to the HEB solution prior to trituration brining the DNase concentration to 0.08 mg/mL, and (2) pelleted dissociated cells are resuspended in rat glia culture media (see recipe in Materials section above). Glial cells are cultured for at least 10 days until the cells have formed a confluent monolayer. Once glial cells reach 100% confluency, they are ready for dissociation and cryobanking. 1. For dissociating rat glial cells, pre-warm 0.25% trypsin-EDTA and rat glia culture medium to 37  C.

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2. Aspirate and completely remove the glial medium from the T-175 flasks, and rinse twice with 60 mL PBS (see Note 7). 3. Aspirate PBS and add 20 mL 0.25% trypsin-EDTA solution. 4. After 10 min at 37  C, gently tap the side of the flask to dislodge the glial cells from the surface. 5. After most cells have lifted, quench the trypsin by adding 20 mL rat glia culture medium to the flask. Pipette up and down 10–18 times with a 10 mL serological pipette to completely detach cells, and transfer the cell suspension to three 15 mL conical tubes per flask (see Note 8). 6. Centrifuge cell suspension for 5 min at 300
g at room temperature (RT) and aspirate the supernatant. Resuspend the cells in 1 mL of rat glia culture medium by gently pipetting up and down three to five times with a P-1000 μL pipettor. 7. To generate cryo-stocks, count the cells in the suspension, and dilute to a concentration of 4–20 million cells/mL (2
the storage concentration) in rat glia culture medium. At a 1:1 volume ratio, add freezing medium to the cell suspension in a dropwise manner, bringing the final volume ratio to 10% DMSO/40% FBS/50% rat glia culture medium. Aliquot this new suspension in 2 mL cryovials with 1 mL/vial for final concentration of 2–10 million cells per vial. The final concentration of cells can be adjusted depending on anticipated downstream use. 8. Immediately after aliquoting, place cryovials in a ~1  C/min isopropyl alcohol freezing container, and store in a 80  C freezer for 16–24 h. 9. Transfer cryovials to liquid nitrogen storage unit for long-term storage. We have used glia cryo-stocks that were preserved for more than 2 years without an obvious decline in cell viability or performance. 4.2.2 Coating of 96-Well Ibidi Plates

1. Coat the 96-well Ibidi plates with poly-D-lysine (PDL) by adding 200 μL of 100 μg/mL PDL solution onto the center of each well (see Note 9). 2. Incubate plates overnight at 4  C before plating neurons.

4.2.3 Dissociation of Rat Hippocampal Tissue

Rat primary hippocampal tissues shipped from BrainBits are stored at 4  C in HEB solution prior to use. Cell viability is high if dissociation occurs within 1–2 days of tissue dissection. 1. Prepare a 15 mL conical tube with the dissociation solution by dissolving sterile papain enzyme in the Hibernate E minus calcium minus B27 solution for a final working concentration of 2 mg/mL. This cell dissociation solution is incubated for 10 min in the 37  C water bath prior to use.

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2. Carefully transfer HEB solution surrounding the tissue to another sterile conical tube for later use. 3. Add 2 mL of cell dissociation solution (papain enzyme solution) to the tissue, and transfer the tissue and solution to a 15 mL conical tube. Incubate for 10 min in the 37  C water bath and gently swirl after 5 min (see Note 10). 4. Remove the cell dissociation solution leaving the tissue at the bottom. Add the HEB solution retained from step 2 to the conical tube with the tissue, and triturate tissue with a 1000 μL pipette tip for 1–2 min while carefully avoiding air bubbles. Let the undispersed tissue pieces settle for 1 min. 5. Transfer supernatant containing dissociated cells to a sterile 15 mL conical tube, discarding the last ~50 μL of HEB solution containing large tissue pieces and other debris. Pellet the cells by centrifuging for 2 min at 300
g at RT, and discard the supernatant leaving behind ~100 μL of HEB solution containing the cell pellet. Disperse the pellet of cells by flicking the bottom of the tube, and then resuspend in 1 mL of rat neuronal culture medium (see recipe in Materials section above). 4.2.4 Plating of Hippocampal Neurons and Glial Cells

1. Thaw required number of stored glia cryovials, and count the cells with a hemocytometer or automated cell counter. 2. In rat neuronal culture medium at RT, prepare a mixed cell suspension of rat hippocampal neurons and glial cells. The required number of cells and volume per well is 12,000 neurons and 15,000 glial cells in a 500 μL volume. Since surface area per well of an Ibidi 96-well plate is 0.56 cm2, the final density of plated cells is ~21.4
103 neurons/cm2 and ~ 26.8
103 glial cells/cm2. 3. Prior to plating, equilibrate the PDL-coated plates to RT (since the plates are stored at 4  C after addition of PDL solution). Aspirate the PDL solution from the PDL-coated 96-well Ibidi plates, and rinse each well once with 200 μL Neurobasal-A medium. 4. Plate 500 μL of the neuronal and glial cell suspension onto each well. Incubate the plates at RT for 30 min before transferring to a 37  C, 5% CO2 incubator (see Note 11). 5. Cells are maintained in the incubator without any interventions until DIV6, when they are ready for lentiviral delivery of constructs encoding CheRiff and jRGECO1a.

4.2.5 Lentiviral Transduction and Additional Culture

Rat neuronal cultures are treated with lentiviral particles encoding the optogenetic and calcium sensor constructs 6 days after plating (DIV6), at which point neurons have firmly attached to the substrate and developed axons and dendrites (Fig. 8a).

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Fig. 8 Rat hippocampal cultures. Primary E18 rat hippocampal neurons co-cultured with rat glial cells and plated at ~21,400 cells/cm2 on Ibidi 96-well plates. (a) Phase contrast image 6 days after plating and prior to lentiviral transduction. Initial neuronal processes are clearly visible. (b) 12 days after plating and 6 days after transduction, more extensive neuronal processes are visible

1. Warm frozen lentiviral stocks to RT, and add them to rat neuronal culture medium with a multiplicity of infection (MOI) ratio of 2 parts CheRiff-EBFP2 per 1 part Syp-jRGECO1a or 4 parts CreON-jRGECO1a, bringing the total well volume to 200 μL. For our standard 30x-concentrated lentivirus and typical MOIs, this corresponds to 0.3 μL per well of CheRiff-encoding lentiviral particles and 0.15 μL per well of Syp-jRGECO1a-expressing lentivirus. 0.01 μL of Cre balances the number of CheRiff- and jRGECO1a-expressing neurons in the synaptic transmission assay (Fig. 6c) (see Note 12). 2. Incubate neuronal cultures with the virus for at least 16 h (inside the 37  C, 5% CO2 incubator). 3. At DIV7, aspirate the medium containing the virus from the wells, leaving behind ~50 μL, and add 400 μL of fresh rat neuronal culture medium to each well (see Note 13). 4. At DIV12 before the plates are functionally imaged, aspirate ~350 μL of rat neuronal culture medium, and replace with 400 μL of fresh medium. By this point, small numbers of neurons may clump together, but most should remain well dispersed with extensive neuronal processes visible by phasecontrast imaging (Fig. 8b). Transduction efficiency should be high, with more than ~90% of neurons expressing the optogenetic constructs. The pre-synaptic calcium sensor should show punctate expression (Fig. 2b), and CheRiff should be membrane localized (Fig. 2c). For the synaptic transmission assay, the jRGECO1a and the CheRiff should be expressed in minimally overlapping sets of neurons (Fig. 5e).

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In order to measure synaptic signaling in primary rat neuronal cultures, we functionally image the cells using a custom ultrawidefield microscope known as Firefly [16]. A 570 nm yellow laser is used to excite the jRGECO1a calcium sensor fluorescence, and a 470 nm blue LED, patterned by a digital micromirror device (DMD), is used to stimulate CheRiff. 1. Prior to imaging, the neuronal culture media is exchanged with Tyrode’s imaging buffer (see recipe in Materials section above) to reduce background autofluorescence while maintaining stable cellular physiology at ambient CO2 levels. Remove the 96-well Ibidi plate from the incubator, aspirate culture media, and gently wash twice with 300 μL per well of Tyrode’s buffer. Fill the well with 300 μL Tyrode’s buffer, and incubate for at least 15 min at RT to allow cells to equilibrate to their new temperature and buffer. 2. Wipe the bottom of the 96-well plate with a clean dry Kimwipes to remove any dust or dirt before placing onto the microscope’s automated translation stage (see Note 14). 3. A 570 nm yellow laser with an intensity of 10–135 mW/cm2 is used to excite the jRGECO1a calcium sensor and record calcium signaling. At an intensity of 135 mW/cm2, the yellow laser-induced current through CheRiff is minimal [16]. 4. Another type of crosstalk occurs when the blue light that is used to stimulate CheRiff drives the jRGECO1a calcium sensor into a brighter photophysical state [14, 15, 39]. To avoid this crosstalk, a digital micromirror device (DMD) is used to pattern the 470 nm blue light so no blue light illuminates the record region (see Note 15). 5. An “Oreo” pattern is used to stimulate outside the field of view (FOV) where the calcium signal is recorded (Fig. 1). Movies are recorded at 100 Hz. The calcium recording region fills the 1 mm gap between the blue stimulus regions (see Note 16). 6. Motorized stages are used to image three FOVs per well. Ensure that the FOVs are appropriately spaced as to not stimulate the same cells more than once. 7. The stimulus protocol used consists of five identical 5 ms pulses that are spaced 3 s apart. The blue light intensity used was 125 mW/cm2, which reliably elicits action potentials in CheRiff-expressing neurons. Example recordings for both assays are shown in Figs. 4 and 6.

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Notes 1. 293T cells are passaged when they reach 80–100% confluency. After the thaw, cells divide slowly, so we use a ~1:1 dilution ratio (half a 15 cm plate (frozen) onto one 10 cm plate). After the first passage, cells are still not dividing at full speed, so we use a ~1:2 dilution ratio (one 10 cm plate into one 15 cm plate). After that, 293T cells typically double once per day, and they can be passaged every other day at a 1:4 ratio or every third day at a 1:8 ratio. 2. Each HYPERFlask can hold around 560 mL volume. Refer to the online video from Corning Life Sciences [40] for a practical demonstration. 3. To ensure efficient transfection, use endotoxin-free plasmid DNA prepared using a QIAGEN Maxi scale or larger prep. The DNA plasmid should be at a concentration ≳1500 ng/μL. 4. We have observed that medium containing antibiotics (P/S) may reduce the yield of lentiviral particles. Therefore, for this step we recommend using medium without antibiotics (Lentiviral Production Medium 1). 5. We have observed no adverse effects in the functional performance of neurons from the Lenti-X Concentrator reagent. 6. Although we have not quantified lifetime, we have successfully used lentiviral particles stored at 80  C virus for at least 2 years. However, freeze/thaw cycles should be avoided as these lower viral titers. 7. Washing the T-175 flasks twice with PBS ensures complete removal of serum-containing rat glial culture medium, which can quench trypsin activity. 8. Do not incubate the cells with trypsin longer than 10 min, as it may adversely affect cell health and viability. Proceed with the protocol even if some cells remain attached to the surface of the flask after pipetting up and down 15–20 times with the serological pipette. 9. For some neuronal types, a combination of PDL and recombinant laminin coatings provides higher viability, but for rat hippocampal neurons, we have found that the simpler and less expensive PDL coating alone resulted in equal performance. 10. While swirling, the two tissue pieces should begin to adhere to each other, indicating that tissue digestion has progressed sufficiently to begin trituration. 11. Having both media and plates at RT eliminates convection currents during the initial 30 min incubation at RT, allowing cells to settle directly to the bottom surface of the well and

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attach with a uniform density. By the time the plates are transferred into the 37  C, 5% CO2 incubator, cells have attached, and convection currents as the plates heat up have minimal impact on cell distribution. 12. A reduced volume per well is used during transduction to increase viral concentration and infection efficiency. For CheRiff, we have produced three HYPERFlask batches with concentrations ranging from 4
1010 to 6
1010 genome copies/ mL. In a typical transduction, we use an in-well virus volume fraction of 0.17% with a stock virus concentration 5
1010 genome copies/mL. We explored whether transduction efficiency depends on virion concentration, which would be expected if the virus concentration is minimally reduced by cell infection; virion number per area, which would be expected if the virus sinks to the bottom of the well; or multiplicity of infection (MOI), which would be expected if most of the added virions successfully infect a cell. We found that transduction efficiency depends on all three, so experimental conditions do not fall cleanly into a single limiting case. As a result, ideally, all three parameters should be held constant when switching plate format. Typical values we use for CheRiff are as follows: virus concentration, 6
107 genome copies/mL; virion number per area, 3
107 genome copies/cm2; and MOI, 1
103 genome copies/plated neuron. The fraction of viable, infectious virions can differ markedly from prep-to-prep, and a functional virus titration of each round is required to achieve optimal assay performance. 13. Avoid aspirating the entire volume of medium from the well, which increases the risks of cell drying or disruption from strong flow forces during medium re-addition. In our experience, the residual virus has minimal adverse effects on neuronal health. 14. The published Firefly microscope [16] uses a prism just below the sample to couple the laser into the cells at a near-total internal reflection (TIR) angle where the laser propagates into the imaging buffer nearly parallel to the sample surface. For voltage imaging with Optopatch [13, 16], this is important to increase the red laser intensity and reduce background autofluorescence because the voltage sensor QuasAr is very dim. When imaging the much brighter calcium sensor jRGECO1a, through-prism near-TIR illumination adds complexity but minimally improves the signal-to-noise ratio (SNR), so the prism may be omitted. When imaging with the prism, apply four to five drops of microscope oil on the center of the prism, and carefully place 96-well Ibidi plate on top of the prism to ensure that there are no air bubbles that can distort the image.

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15. For the synaptic transmission assay (Fig. 5), two types of crosstalk are suppressed. The patterned illumination prevents optical crosstalk where the blue light perturbs jRGECO1a. Expressing the sensors in non-overlapping populations of neurons ensures that the blue stimulus will not excite the neurites of any cells that extend from cell bodies in the recording region out into the stimulus region. This combination ensures that the great majority of the observed signal is mediated by synaptic transmission—the detected signal is almost completely eliminated by the addition of synaptic blockers (Fig. 6a, b). 16. In order to obtain robust signals in the pre-synaptic calcium assay, it is essential that axons from neurons in the optogenetically stimulated region (blue regions in Fig. 1b) extend throughout the recording region. For the synaptic transmission assay, the axons from neurons in the stimulated region must extend into the recording region, and/or the dendrites from neurons in the recording region must extend into the stimulated region. To measure the characteristic neurite length scale, we used the DMD to stimulate a 500-μm-wide vertical stipe in the pre-synaptic calcium assay and measured the signal in 250-μm-wide vertical stripes moving away from the stimulus region. The signal showed approximately exponential decay as with a half-length of ~600 μm, which approximates the axon extent. The center of the 1 mm gap in stimulation is 500 μm from stimulated area above and below and displays robust calcium signaling in both assays. References 1. Saitsu H, Kato M, Mizuguchi T et al (2008) De novo mutations in the gene encoding STXBP1 (MUNC18-1) cause early infantile epileptic encephalopathy. Nat Genet 40:782–788. https://doi.org/10.1038/ng.150 2. Abramov E, Dolev I, Fogel H et al (2009) Amyloid-beta as a positive endogenous regulator of release probability at hippocampal synapses. Nat Neurosci 12:1567–1576. https://doi.org/10.1038/nn.2433 3. Nemani VM, Lu W, Berge V et al (2010) Increased expression of alpha-synuclein reduces neurotransmitter release by inhibiting synaptic vesicle reclustering after endocytosis. Neuron 65:66–79. https://doi.org/10.1016/ j.neuron.2009.12.023 4. Durand CM, Betancur C, Boeckers TM et al (2007) Mutations in the gene encoding the synaptic scaffolding protein SHANK3 are associated with autism spectrum disorders. Nat Genet 39:25–27. https://doi.org/10.1038/ ng1933

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Chapter 9 Optogenetics to Interrogate Neuron-Glia Interactions in Pups and Adults Chloe´ Habermacher, Blandine Manot-Saillet, Domiziana Ortolani, Fernando C. Ortiz, and Marı´a Cecilia Angulo Abstract In just over 10 years, the use of optogenetic technologies in neuroscience has become widespread, having today a tremendous impact on our understanding of brain function. An extensive number of studies have implemented a variety of tools allowing for the manipulation of neurons with light, including lightactivated ion channels or G protein-coupled receptors, among other innovations. In this context, the proper calibration of photostimulation in vivo remains crucial to dissect brain circuitry or investigate the effect of neuronal activity on specific subpopulations of neurons and glia. Depending on the scientific question, the design of specific stimulation protocols must consider from the choice of the animal model to the light stimulation pattern to be delivered. In this chapter, we describe a detailed framework to investigate neuron-glia interactions in both mouse pups and adults using an optogenetic approach. Key words Optogenetics, Channelrhodopsin, Patch-clamp recordings, Local field potentials, Immunostainings, Neuron-glia interactions, Oligodendrocytes

1

Introduction Using light to control neurons expressing opsin-derived proteins ensures the manipulation of their activity over different timescales and spatial precision. A major advantage of optogenetics is the possibility to target and manipulate specific cell types among brain regions by the expression of light-sensitive ion channels, transporters, or receptors. Two versatile approaches enable the expression of these proteins in neurons: virus injection (e.g., AAV or lentivirus vectors) and transgenic animals (e.g., Cre-Lox system). However, the efficiency of expression may differ between brain structures, developmental stages, and gene expression promoters [1, 2]. Indeed, to achieve a stable expression over time remains a major limitation. Given that a central parameter affecting optogenetic control is the level of light-sensitive protein expression, scientists must carefully characterize their model considering that

Robert E. Dempski (ed.), Channelrhodopsin: Methods and Protocols, Methods in Molecular Biology, vol. 2191, https://doi.org/10.1007/978-1-0716-0830-2_9, © Springer Science+Business Media, LLC, part of Springer Nature 2021

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over-expression may induce a block of the depolarization or even turned to be toxic [3]. Since the intrinsic membrane properties of neurons may vary from one subtype to another, an induced photocurrent can be differently translated into membrane potential changes and, therefore, have distinct consequences on neuronal network function. Moreover, the properties of the opsin, such as the precise activation wavelength, kinetics of the response, and dependency on other factors (i.e., voltage or temperature), need careful consideration. Therefore, to validate a particular photostimulation protocol in inducing neuronal and network responses, it is crucial to test it in ex vivo tissue, such as acute brain slices, before performing in vivo experiments. The frequency of stimulation must be consistent with the level of opsin expression and firing frequency of the stimulated neuronal cell type, e.g., fast-spiking interneurons or pyramidal cells in the cerebral cortex [2, 4]. This step should include the evaluation of (i) the functional expression of the light-sensitive protein, (ii) the ability of the targeted cell to respond to a light-train stimulation protocol (i.e., whether or not it is able to follow the simulation frequency, among other parameters), and (iii) a synchronous activation of a neuronal population, for instance, by recording lightevoked local field potentials (LFPs). Then, the calibration of in vivo experiments is crucial to ensure that the pattern of delivered light is sufficient to trigger an efficient repeated stimulation. One strategy consists in coupling the optical fiber with a recorded electrode (called optrode) to monitor the effect of light pulses on neural activity. Finally, photostimulation can also induce relevant increases in the temperature of the tissue [5–7], an often overlooked effect that can modify cell proliferation and even the gating properties of the opsin itself [7–10]. Thus, control experiments using wild-type mice lacking the light-sensitive protein are also important. In this chapter, we describe a detailed procedure using transgenic mice expressing channelrhodopsin-2 (ChR2) fused with eYFP to obtain an efficient in vivo photostimulation of specific neurons in pups and adults and interrogate the effect of neuronal activity on oligodendroglia dynamics [2, 4, 11].

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Materials

2.1 Verification of Opsin Protein Expression in Target Neurons

Prepare all solutions using ultrapure water (prepared by purifying deionized water to attain a resistivity of 18 MΩ cm). 1. Phosphate-buffered saline (PBS) made with ultrapure water. 2. Solution for anesthesia: mix of ketamine (20 mg/mL)/xylazine (2 mg/mL) in filtered PBS (adjust to a final dose of 0.1/ 0.01 mg/g ketamine/xylazine).

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3. Phosphate buffer 0.15 M containing 4% paraformaldehyde, pH 7.4. 4. Dissection material (scissors, spatula, forceps). 5. Blades. 6. Glue (ethyl-2-cyanoacrylate). 7. Filter paper. 8. Vibratome. 9. Orbital shaker. 10. Permeabilization solution: 1% Triton X-100, 4% normal goat serum (NGS). Dilute the appropriate volume of NGS in PBS and add Triton drop by drop (see Note 1). 11. Incubation solution: 0.2% Triton X-100, 2% NGS. Dilute the appropriate volume of NGS in PBS, and add the Triton drop by drop (see Note 1). Then, add the appropriate volume of specific antibodies labelling the target neurons (e.g., Parvalbumin for fast-spiking cells in the cortex). If the expression of ChR2 fused to YFP is high, it should not be necessary to amplify the YFP signal. 12. Glass slides and coverslips. 13. Mounting medium. 14. Confocal microscope. 2.2 Implementation of the Photostimulation Protocol in Acute Brain Slices 2.2.1 Preparation of Acute Brain Slices

1. Dissection material (scissors, spatula, forceps). 2. Blades. 3. Glue (ethyl-2-cyanoacrylate). 4. Filter paper. 5. Vibratome. 6. Water bath. 7. Cutting solution for pups: 215 mM sucrose, 2.5 mM KCl, 1.25 mM NaH2PO4, 26 mM NaHCO3, 20 mM glucose, 5 mM pyruvate, 1 mM CaCl2, 7 mM MgCl2, pH 7.35. Dilute in a glass beaker the amount of sucrose, KCl, NaH2PO4, NaHCO3, glucose, and pyruvate required for a final volume of 500 mL in 400 mL of ultrapure water (see Note 2). Bubble this solution with a 95% O2/5% CO2 gas mix during 10 min, transfer the solution in a 500 mL volumetric flask, add the required volume of CaCl2 and MgCl2 solutions (see Note 3), and adjust to the final volume. 8. Cutting solution for adult mice: 93 mM N-methyl-D-glucamine (NMDG), 2.5 mM KCl, 1.25 mM NaH2PO4, 30 mM NaHCO3, 20 mM HEPES, 2 mM thiourea, 25 mM glucose, 5 mM ascorbate, 3 mM pyruvate, 0.5 mM CaCl2, 10 mM MgCl2, pH 7.35.

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Dilute in a glass beaker the required amount of NMDG, KCl, NaH2PO4, NaHCO3, HEPES, thiourea, glucose, ascorbate, and pyruvate for a final volume of 500 mL, in 400 mL of ultrapure water (see Note 4). Adjust the pH at 7.4 by carefully adding (drop by drop) a 10 M HCl solution. Add the required volume of CaCl2 and MgCl2 solutions (see Note 3), transfer the solution in a 500 mL volumetric flask, and adjust the volume to the final volume. 9. Perfusion solution: 126 mM NaCl, 2.5 mM KCl, 1.25 mM NaH2PO4, 26 mM NaHCO3, 20 mM glucose, 5 mM pyruvate, 2 mM CaCl2, 1 mM MgCl2, pH 7.4. Dilute in a glass beaker the appropriate quantity of NaCl, KCl, NaH2PO4, NaHCO3, glucose, and pyruvate required for a final volume of 1 L, in 800 mL of ultrapure water (see Note 5). Bubble this solution with a 95% O2/5% CO2 gas mix during 10 min, transfer the solution in a 1 L volumetric flask, add the required volume of CaCl2 and MgCl2 solutions (see Note 3), and adjust to the final volume. 2.2.2 Electrophysiological Recordings in Whole-Cell Configuration During Photostimulation in a Patch-Clamp Rig

1. Intracellular (intrapipette) solution: 130 mM K-gluconate, 0.1 mM EGTA, 0.5 mM CaCl2, 2 mM MgCl2, 2 mM Na2ATP, 0.5 mM Na-GTP, 10 mM HEPES, 10 mM phosphocreatine. Dilute K-gluconate and EGTA in 35 mL of ultrapure water, adjust the pH at 7.4 with a solution of KOH, add the required volumes of CaCl2 and MgCl2 solutions and dissolve HEPES, and adjust the pH at 7.4. Put the solution on ice while mixing. Dissolve Na2-ATP, Na-GTP, and phosphocreatine. Adjust the pH at 7.4. Complete the volume to 50 mL using a volumetric flask. Filter the solution through 0.22 μm membrane. Measure the osmolarity of the solution (290–310 mOsm), make 1 mL aliquots, and store at
20  C. 2. Epifluorescence microscope equipped with a perfusion chamber. 3. Perfusion system. 4. Patch-clamp amplifier. 5. Headstage. 6. Puller. 7. Patch pipettes: set a specific program on the puller to obtain a resistance of 3–5 MΩ from glass capillaries (1.2 OD  0.69 ID). 8. Recording electrode. 9. Light source for photostimulation: ultra-high-power LED system able to deliver the desired wavelength. 10. Current controller to feed the light source.

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11. Optic fibers. 12. Pulse generator device to create programmable TTL pulses from a software. 2.3 Surgical Procedure

Prepare all injecting solutions using phosphate-buffered saline (PBS) made with ultrapure water (prepared by purifying deionized water to attain a resistivity of 18 MΩ·cm) filtered through a 0.22 μm membrane. 1. Buprenorphine solution (0.01 mg/mL). 2. Optical power meter. 3. Solution for anesthesia: mix of ketamine (20 mg/mL)/xylazine (2 mg/mL) in filtered PBS (adjust to a final dose of 0.1/ 0.01 mg/g ketamine/xylazine). 4. Razor. 5. Stereotaxic frame. 6. Ophthalmic dexpanthenol gel. 7. Lidocaine solution 1%. 8. Skin disinfectant solution and cotton swab. 9. Surgical instruments: scissors, blades, scalpel, forceps. 10. Stereotaxic frame for mice. 11. HCl solution (1 mM). 12. Glue (ethyl-2-cyanoacrylate). 13. Cannula accommodating a mini-optic 1.25 mm; fiber, 200 μm diameter).

fiber

(cannula,

14. Dental cement including a kit to mix it. 15. Suture material. 2.4 Control of the Efficiency of Photostimulation In Vivo

1. All the material described in Subheading 2.3. 2. Optrode: Cannula containing a mini-optic fiber surrounded by a 50 μm wire electrode (Ni/Ag). The tip of the wire must be placed just below the tip of the mini-optic fiber and attached to a pin connector (see Note 6). 3. 50 μm wire electrode (Ni/Ag) attached to a pin connector to be used as a ground. 4. Two fiber optic patch cords. 5. Rotary joint. 6. Material described in Subheading 2.2.2 from items 2 to 5 (electrophysiological setup). 7. Headstage of the amplifier. 8. Custom-made Faraday cage (see Note 7).

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2.5 In Vivo Stimulation of Awake Mice

1. Material described in Subheading 2.4 from items 4 to 6 (stimulation setup).

2.6 Immunostaining to Evaluate Activity-Dependent Neuron-Glia Interactions

1. Material described in Subheading 2.1 (immunostaining protocol).

2. Open-field arena.

2. Primary antibodies to label oligodendroglia: anti-Olig2 and anti-CC1. Dilute each antibody in the incubation solution at the recommended concentration (1:400 for anti-Olig2 and 1:100 for anti-CC1) (see Note 8). 3. Appropriate secondary antibodies diluted in the incubation solution at 1:500. 4. A stock solution of 5-ethynyl-20 -deoxyuridine (EdU) (10 mM) diluted in filtered PBS stored at
20  C. 5. Commercially available kit for EdU revealing. Clik-iT EdU Alexa Fluor 647R (use according to the manufacturer’s instructions).

3

Methods

3.1 Evaluation of Protein Expression

1. Anesthetize the mouse following guidelines for the care and use of laboratory animals with an intraperitoneal injection of the ketamine/xylazine mix. 2. Perfuse the mouse transcardially, first with PBS (15 mL) and then with a 4% PFA solution (40 mL) at a flow around 3 mL/ min (see Note 9). 3. Extract the brain and put it 2 h at room temperature in a 4% PFA solution. 4. Wash the brain with chilled PBS during 10 min three times, and store it at 4  C (see Note 10). 5. Cut a small brain piece with a blade to make a plane edge surface. 6. Stick the brain with glue on the plate of the vibratome, and cover it with chill (almost frozen) PBS (see Note 11). 7. Cut 50–100-μm-width slices with the vibratome using the appropriate frequency and speed parameters to preserve the tissue. 8. Collect the slices of the region under study by taking them gently with a small paintbrush. Transfer them to a multi-well plate previously filled with chilled PBS. 9. Put slices in the permeabilization solution during 2 h at room temperature and gentle shaking. 10. Wash slices with PBS during 10 min three times.

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11. Incubate slices in the solution containing primary antibodies prepared at recommended dilutions and incubation times (see Note 8). 12. Wash slices with PBS during 10 min three times. 13. Incubate slices in the solution containing secondary antibodies prepared according to the recommended dilutions and incubation times (see Note 12). 14. Mount slices on a slide by manipulating them gently with a small paintbrush. 15. Cover the slices with an appropriate mounting medium (i.e., fluorescence mounting medium). Cover them with a coverslip. 16. Take confocal images of the region of interest. 17. Check that ChR2-expressing cells correspond to the targeted ones by the co-localized expression of the ChR2-reporter protein expression (i.e., YFP) and a specific cell marker. 3.2 Setting the Photostimulation Protocol in Acute Brain Slices 3.2.1 Preparation of Acute Brain Slices

The general procedure to perform experiments in brain slices is illustrated in Fig. 1.

1. Chill the cutting solution (see Note 13), and bubble it with 95% O2/5% CO2 gas mix. Bubble the perfusion solution at room temperature with the same mix. Put an incubation chamber filled with perfusion solution at 34  C (in the water bath) while bubbling it with the same gas mix. 2. For adult mice slicing, put a second incubation chamber with a bubbled cutting solution at 34  C in the water bath. 3. Anesthetize the mouse following guidelines for the care and use of laboratory animals with an intraperitoneal injection of the ketamine/xylazine mix. 4. Kill pups by decapitation or adult mouse by cervical dislocation, cut the head, and put it quickly in a glass beaker containing cold cutting solution. 5. Extract the brain and cut it with a blade to make a plane surface (see Subheading 3.1, step 5). 6. Stick the brain with glue on the plate of the vibratome, and add cold cutting solution. 7. Cut 300-μm-thick slices with the vibratome using the appropriate frequency and speed parameters to preserve the tissue. 8. For pups, put the slices in the incubation chamber containing perfusion solution at 34  C. Once the last slice is cut, wait 15 min, remove the incubation chamber from the bath, and wait 30 min at room temperature.

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Fig. 1 Schematic representation of the procedure for optogenetic stimulation in acute brain slices. (a) Before performing functional experiments, a crucial step is to perform a first evaluation of the opsin protein expression in target cells by using immunostainings at developmental stages pertinent for the study. (b, c). Preparation of acute slices of the brain region under study. Use different procedures in pups and adults (c). (d) Establishment of the photostimulation protocol using patch-clamp recordings of neurons expressing the opsin, e.g., ChR2. Note a decrease in the capacity of neurons to follow the photostimulation train (blue lines) when increasing the light-train frequency from 10 to 50 Hz in pups (d) [2]. (e) Photo-evoked LFPs in acute brain slices. This example shows LFPs (black trace) recorded in cortical layer V of mouse pups during photostimulation (blue lines) with an optic fiber located above the recording site

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9. For adults, put the slices in the incubation chamber containing cutting solution at 34  C (prepared in step 2) during 4–8 min maximum. Then, transfer them to the incubation chamber containing perfusion solution at 34  C. Once the last slice is transferred, wait 15 min, remove the incubation chamber from the bath, and wait 30 min at room temperature. 3.2.2 Electrophysiological Recordings of Individual Neurons During Photostimulation

10. Measure the power at the tip of the optic fiber, and adjust the parameters of the current controller to obtain the desired light power during the experiment (see Note 14). 11. Put a slice in the recording chamber under the microscope of the patch-clamp rig. 12. Patch a cell expressing the reporter (i.e., YFP) fluorescent protein fused to the opsin. 13. Characterize the neuron with a protocol of depolarizing step current injections in current-clamp mode. 14. Put the tip of the optic fiber near the patched cell, apply trains of photostimulation using the desired parameters (i.e., frequency, pulse duration, total time length), and record simultaneously the firing of the patched neuron in currentclamp mode. 15. Test the ability of the patched neuron to sustain its discharge upon current pulse injections mimicking the train of photostimulation used in step 14. 16. Calculate the success rate of action potential discharges during both trains of photostimulation and current pulses at the tested frequencies, and compare them to determine the appropriate frequency generating a minimum of fails during photostimulation. 17. Put an extracellular electrode filled with perfusion solution close to the optic fiber, and record LFPs evoked by light at the frequency determined at step 15.

3.3 Surgical Procedure

The general procedure to perform in vivo experiments is illustrated in Fig. 2. Before starting the surgery, check the parameters of the light delivery setup to ensure the correct outcome power at the tip of the mini-optic fiber. 1. Connect the mini-optic fiber through the cannula to the fiber optic patch cord, ensure the absence of light leakage, measure the power at the tip of the implant with a power meter, and adjust the parameters of the controller to get the correct intensity. 2. Fix the cannula connected to the mini-optic fiber to the holder of the stereotaxic frame.

Fig. 2 Schematic representation of the procedure for optogenetic stimulation in vivo. (a) Stereotaxic surgery to implant the optic fiber in mouse pups (left) and adults (right). The optic fiber within a cannula (Implant) can be placed on the pial surface (left) or inside (right) the brain according to region under study. Skull has to be cleaned from connective tissue and dried. Glue (dark red) is applied in pups to stiffen the skull and in adults to increase the adherence of the cement (dark pink). In adults, flattening the skull helps to fix the optic fiber. (b) Photo-evoked LFPs in an anesthetized mouse using an optrode containing the optic fiber and a recording electrode. This example shows LFPs (black trace) recorded in cortical layer V of mouse pups during photostimulation at the surface of the brain (blue lines) [2]. (c) Photostimulation in freely moving mice without the electrode. To assess proliferation, inject EdU at least 20 min prior to the stimulation. (d) Immunostainings to assess the effect of neuronal activity in oligodendroglia. The mouse is sacrificed after the stimulation, and specific immunostainings are performed to identify the cells (insets). Arrowheads indicate the cells in insets. Scale bars, 20 μm and 10 μm

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3. 30 min before to start the surgery, administer buprenorphine (0.05 mg/kg) by intraperitoneal injection to prevent postprocedural pain. 4. Induce a deep anesthesia (see Note 15), and cover the eyes with eye ointment to prevent dehydration. 5. Shave the head. 6. Fix the mouse in a stereotaxic frame, apply lidocaine on the skin, and disinfect with Betadine. 7. Incise the skin (see Note 16), and clean connective tissue with a blade (see Note 17). 8. For pups, stiffen artificially the skull with two layers of glue (see Note 18). 9. Mark the right position to place the implant according to the desired coordinates, drill carefully the skull without touching the cortical surface of the brain, and remove the residual pieces of bone by using thin forceps. 10. For adult mice, lightly scratch the skull with a small scalpel blade to flat the surface before adding the glue. 11. Clean properly the surrounding area with PBS, and wait to ensure that the skull is dry. 12. For adult mice, gently spread out a drop of glue on the skull around the hole to improve the fixation of the cement. 13. Implant stereotaxically the cannula containing the mini-optic fiber according to the desired coordinates (see Note 19). 14. Apply a first layer of cement with the tip of a curved needle (see Note 20), wait until the cement is almost dry, and apply a second layer of cement (see Note 21). 15. Remove carefully the holder without moving the cannula. 16. Suture the skin around the implant. 3.4 Control of the Efficiency of Photostimulation In Vivo

Implantation of an optrode, containing the cannula, the mini-optic fiber, and the electrode, makes it possible to measure LFPs of neurons in response to photostimulation (see Note 22). 1. Follow the protocol described previously in Subheading 3.3 until step 9. 2. Drill the skull to make a small hole to put the ground electrode (see Note 23). 3. Follow the protocol described in Subheading 3.3 from step 10 to 15. 4. Place the ground electrode at the surface of the brain. Fix it with two layers of cement, and wait until the cement is dry. 5. Suture the skin around the implant and the ground.

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6. Place the anaesthetized mouse inside the custom-made Faraday cage to verify that photostimulation parameters evoked efficient neuron activation (see Note 7). 7. Connect the optic fiber patch cord to the implant, the headstage to the recording electrode, and the ground electrode to the amplifier ground. 8. Set up the parameters on the current controller and the parameters of photostimulation determined previously in Subheading 3.2. 9. Record LFPs evoked by light pulses. 3.5 In Vivo Stimulation of Awake Mice

3.6 Evaluation of Activity-Dependent Neuron-Glia Interactions

1. Connect carefully the implant to the optic fiber patch cord. 2. Set up the parameters on the current controller. 3. Start the protocol of photostimulation controlled by a software generating TTL pulses (see Note 24). In order to evaluate the effect of neuronal activity on oligodendroglia dynamics, EdU assays and specific immunostainings can be performed. 1. 20 min before starting the in vivo stimulation, administer EdU solution (50 mg/kg) by intraperitoneal injection (see Note 25). 2. Repeat procedures described in Subheading 3.5. 3. Repeat procedures described in Subheading 3.1 until step 13. 4. Reveal the EdU-labelled cells by using Clik-iT EdU Alexa Fluor 647R according to the instructions provided by the manufacturer. 5. Mount slices on a slide by manipulating them gently with a small paintbrush. 6. Cover the slices with an appropriate mounting medium (i.e., fluorescence mounting medium). Cover them with a coverslip. 7. Take confocal images of the region of interest (see Note 26). 8. Determine the number of cells expressing EdU and Olig2 and those expressing EdU, Olig2, and CC1 (see Note 27). 9. Standardize the count number by area (or volume) of analyzed regions. Express this value as density to make it comparable. Cells labelled by EdU are proliferating cells labelled in S-phase, cells expressing only Olig2 are considered oligodendrocyte precursor cells (OPCs), and cells expressing both markers (Olig2 and CC1) correspond to mature oligodendrocytes (OLs) (see Note 28).

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Notes 1. Triton can be diluted faster if PBS is warmed to approx. 37  C. Use a bevel cut end micropipette tip to harvest Triton. 2. 10X stock solution of KCl, NaH2PO4, and NaHCO3 can be prepared and stored at 4  C for 1 month to minimize weight errors. The 1X solution can be prepared the day of the experiment by diluting this stock ten times and adding the proper amounts of the remaining components. 3. Use stock solutions of 1 M CaCl2 and 1 M MgCl2. 4. 10X stock solution of KCl, NaH2PO4, NaHCO3, HEPES, and thiourea can be prepared and stored at 4  C for 1 month to minimize weight errors. The 1X solution can be prepared daily by diluting this stock ten times and adding the proper amounts of the remaining components. 5. 10X stock solution of NaCl, KCl, NaH2PO4, and NaHCO3 can be prepared and stored at 4  C for 1 month to minimize weight errors. The 1X solution can be prepared by diluting this stock ten times and adding the proper amounts of the remaining components. 6. To attach the electrode wire to the cannulae containing the mini-optic fiber, use a small piece of adhesive tape, and then cover the joint with dental cement. This way the optrode components will be fixed avoiding their motion during manipulation. 7. Build a small box with aluminum foil. This box must be grounded through a connector and used to cover the mouse during the recording. 8. Although we advise the use of recommended dilutions and incubation times for each antibody, we also suggest checking for protocols of previously published assays. This is a key step that must be considered when multiple proteins are targeted. For instance, in our hands, the anti-CC1 primary antibody had to be incubated for three nights in order to obtain a proper immunostaining of differentiated oligodendrocytes in adult white matter, but just one night was enough for anti-Olig2. 9. If the transcardiac perfusion is working properly at the end of the PBS perfusion, forepaws should be completely pale. This is indicative of blood withdrawal in the superior thoracic cavity. 10. Chill PBS and keep the bottle on ice during the entire procedure. 11. Dry the brain surface that will be in contact with the glue by using a filter paper before putting it on the plate. 12. From this step, protect slices from light.

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13. Some ice crystals are expected in cold solutions. 14. Most common optogenetic protocols deliver light intensity in a range of 0.5–10 mW/mm2 at the tip of the fiber. 15. Choose the anesthesia method (inhaled isoflurane or ketamine/xylazine intraperitoneal injection (0.1/0.01 mg/g)) according to the target brain area and duration of the surgery. The mask delivering isoflurane could limit the access to place the implant. 16. Make a large incision, and clear the skin to avoid any disturbance during the cementing of the implant. 17. On pups, the conjunctive tissue could be removed by the application of a small drop of HCl solution (1 mM) on a restricted area avoiding any contact with other tissue to improve the adherence of the glue. 18. A first layer of glue is obtained by gently spreading out a drop on the skull. After 10 min, apply a second drop, and wait 20 min to ensure that the glue is completely dried. 19. Descend the optic fiber until touching the surface of the brain. If deep layers are targeted, use optical fibers as thin as possible, and penetrate quickly into the brain to reduce mechanical damage of brain tissue. 20. Prepare the cement mix just before its application and wait for its correct consistency. 21. Apply a first layer of cement to make the foundation and, then, a second one to add strength. 22. Repeat this procedure in control wild-type mice (not expressing the light-sensitive protein). This is central to exclude any optical artifact or heating effect. 23. Place the ground electrode on the skull surface as far as possible from the optrode. 24. Monitor the mouse to ensure that photostimulation does not induce epileptic seizures. If it happens, increase the time between two trains of photostimulation, and/or decrease the power of light (need to restart Subheading 3.4). 25. After the EdU injection, wait at least 20 min before beginning the stimulation to allow EdU to diffuse in the brain. One or two injections (before and at the end of the stimulation) reveal the fate of the cells. Proceed to repeated injections according to the protocol of stimulation and/or the aim of the study. 26. In order to carry on co-localization analyses, the acquisition parameters must consider a proper Z-axis distance and overlapping in between the acquired planes. 27. Count cells in an appropriate software (e.g., ImageJ, free license).

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28. This procedure must be repeated in photostimulated control wild-type mice (with no opsin expression) in order to discard any unspecific effect (e.g., heating).

Acknowledgments This work was supported by grants from Fondation pour la Recherche Me´dicale (FRM; “Equipe FRM DEQ20150331681”), Fondation pour l’aide a` la recherche sur la scle´rose en plaques (ARSEP), a subaward agreement from the University of Connecticut with funds provided by Grant No. RG-1612-26501 from the National Multiple Sclerosis Society (NMSS) to M.C.A. and from ECOS-Sud (Project No. ECOS180013) to M.C.A. and F.C.O. Its contents are solely the responsibility of the authors and do not necessarily represent the official views of UConn or NMSS. F.C.O. was recipient of a FRM post-doctoral fellowship and is supported by the Chilean National Fund for Scientific and Technological Development (FONDECYT, INICIACION #11160616). F.C.O. and C.H. were recipients of ARSEP post-doctoral fellowships. B.M.S. was supported by a PhD fellowship from the French Ministry of Research (ED BioSPC), and D.O. was supported by a INSPIRE PhD fellowship from Marie Skłodowska-Curie Action (European Union’s Horizon 2020) and Universite´ de Paris (grant agreement: 665850; ED BioSPC). References 1. Bitzenhofer SH, Ahlbeck J, Hanganu-Opatz IL (2017) Methodological approach for optogenetic manipulation of neonatal neuronal networks. Front Cell Neurosci 11:239 2. Ortolani D, Manot-Saillet B, Orduz D et al (2018) In vivo optogenetic approach to study neuron-oligodendroglia interactions in mouse pups. Front Cell Neurosci 12:477 3. Lin JY (2011) A user’s guide to channelrhodopsin variants: features, limitations and future developments. Exp Physiol 96:19–25 4. Ortiz FC, Habermacher C, Graciarena M et al (2019) Neuronal activity in vivo enhances functional myelin repair. JCI Insight 4(9): e123434 5. Rungta RL, Osmanski B-F, Boido D et al (2017) Light controls cerebral blood flow in naive animals. Nat Commun 8:14191 6. Senova S, Poupon C, Dauguet J et al (2018) Optogenetic Tractography for anatomofunctional characterization of cortico-

subcortical neural circuits in non-human primates. Sci Rep 8(1):3362 7. Chater TE, Henley JM, Brown JT et al (2010) Voltage- and temperature-dependent gating of heterologously expressed channelrhodopsin-2. J Neurosci Methods 193:7–13 8. Hossain ME, Matsuzaki K, Katakura M et al (2017) Direct exposure to mild heat promotes proliferation and neuronal differentiation of neural stem/progenitor cells in vitro. PLoS One 12(12):e0190356 9. Laissue PP, Alghamdi RA, Tomancak P et al (2017) Assessing phototoxicity in live fluorescence imaging. Nat Methods 14:657–661 10. Stujenske JM, Spellman T, Gordon JA (2015) Modeling the spatiotemporal dynamics of light and heat propagation for in vivo optogenetics. Cell Rep 12:525–534 11. Gibson EM, Purger D, Mount CW et al (2014) Neuronal activity promotes oligodendrogenesis and adaptive myelination in the mammalian brain. Science 344:1252304

Chapter 10 Chronic Optogenetic Pacing of Human-Induced Pluripotent Stem Cell-Derived Engineered Cardiac Tissues Marc Dwenger, William J. Kowalski, Hidetoshi Masumoto, Takeichiro Nakane, and Bradley B. Keller Abstract The delivery of cells into damaged myocardium induces limited cardiac regeneration due to extensive cell death. In an effort to limit cell death, our lab formulates three-dimensional matrices as a delivery system for cell therapy. Our primary work has been focused on the formation of engineered cardiac tissues (ECTs) from human-induced pluripotent stem cell-derived engineered cardiac cells. However, ECT immaturity hinders ability to fully recover damaged myocardium. Various conditioning regimens such as mechanical stretch and/or electric pacing have been used to activate maturation pathways. To improve ECT maturity, we use non-contacting chronic light stimulation using heterologously expressed light-sensitive channelrhodopsin ion channels. We transduce ECTs with an AAV packaged channelrhodopsin and chronically optically pace (C-OP) ECTs for 1 week above the intrinsic beat rate, resulting in increased ECT electrophysiological properties. Key words Cardiac tissue engineering, Optogenetics, Cell therapy, Cardiovascular regeneration, Cell maturation

1

Introduction Higher vertebrates are unable to regenerate cardiomyocytes lost to trauma, leading to pump dysfunction and heart failure [1, 2]. Numerous cell therapy strategies attempt to overcome this regenerative limitation through transplantation of healthy cells into damaged tissue [2, 3]. However, cells used for cell therapy have a very low retention rate when injected into damaged myocardium [3]. The incorporation of cells within biomaterials via tissue engineering increases cell retention and improves differentiation [4]. Our lab developed protocols to construct engineered cardiac tissues (ECTs) from the mixture of cardiac cells, liquid type I collagen, Matrigel, and growth supplements using a vacuumsuctioned mold [5]. However, ECTs remain relatively immature [6]. The constructs possess immature sarcomeres, having reduced

Robert E. Dempski (ed.), Channelrhodopsin: Methods and Protocols, Methods in Molecular Biology, vol. 2191, https://doi.org/10.1007/978-1-0716-0830-2_10, © Springer Science+Business Media, LLC, part of Springer Nature 2021

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force production, and exhibit spontaneous beating [6]. Bioreactors can improve ECT maturity by using mechanical stretch, medium perfusion, and electrical stimulation [6, 7]. However, these approaches require complex parallel support systems, and in particular, electrical stimulation can be invasive, produces Faradaic reactions, and has low spatiotemporal resolution [8–12]. An alternative to electrical stimulation is the optical stimulation of heterologously expressed light-sensitive ion channels, channelrhodopsins, to produce action potentials [13]. Several cardiac optogenetic studies demonstrate the vast capabilities of this technique [12, 14– 17]. To corroborate and extend previous studies, as outlined in the protocol below, we developed a chronic optogenetic ECT conditioning protocol. We expressed a channelrhodopsin variant, ChIEF, in ECTs composed of cardiomyocytes (CM), endothelial cells (EC), and mural cells (MC), all derived from human-induced pluripotent stem cells (h-iPSCs). We cultured ECTs for 7 days unstimulated, followed by 7 days of chronic optical pacing (C-OP). We then assessed ECT maturity by mechanical force testing, histology, RT-qPCR, and immunohistochemistry. This methods chapter will detail protocols involving differentiation of h-iPSCs into various cardiovascular lineages, C-OP, and force testing of ECTs.

2

Materials

2.1 Differentiation of Human iPSCs into Cardiac Cells

1. h-iPSC cell line, undifferentiated with four factors (Oct3/4, Sox2, Klf4, and c-Myc) (see Note 1). 2. Mouse embryonic fibroblast (MEF) cells (see Note 2). 3. Phosphate-buffered saline (PBS) without Ca2+ and Mg2+. 4. High glucose Dulbecco’s Modified Eagle Medium (DMEM) (purchased premade, Life Technologies): Store at 4
C. 5. RPMI 1640 medium (purchased premade from Life Technologies): Store at 4
C for 12 months. 6. Knockout Dulbecco’s Modified Eagle Medium (DMEM) (purchased premade from ThermoFisher Scientific): Store at 4
C. 7. Fetal bovine serum (FBS): Store at 20
C (see Note 3). 8. Fetal calf serum (FCS): Store at 20
C (see Note 3). 9. Knockout serum replacement (KSR) (purchased premade from ThermoFisher Scientific). Store at 20
C (see Note 3). 10. L-Glutamine. 11. 100 nonessential amino acids. 12. B27 supplement without insulin: Store at 20 12 months (see Note 3).




C for

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13. Tissue culture grade 2-mercaptoethanol: Store at 4
C. 14. Versene: Store at 4
C for 18 months. 15. Bovine serum albumin: Store at 4
C. 16. Mitomycin C: Store at 4
C. Reconstitute in sterile water and store at 4
C. Use within 2 weeks of reconstituting. 17. Growth factor-reduced Matrigel: Store 100 μL aliquots at 20
C. Refer to Matrigel lot information for expiration date. 18. Thin-coat Matrigel: Dilute growth factor-reduced Matrigel with 4
C DMEM to a ratio of 1:60 (see Note 4). 19. Human basic fibroblast growth factor (hbFGF): Dilute in PBS or distilled water such that the working solution is appropriate to dilute to 1–10 ng/mL in cell media. Store aliquots of working solution at 20
C for 3 months (see Note 3). 20. Activin A: Reconstitute in 4 mM HCl and aliquot. Concentration should be appropriate to dilute to 100 ng/mL in cell media. Store aliquots at 80
C for 3 months (see Note 3). 21. Wnt3a: Reconstitute in PBS supplemented with 0.1% bovine serum albumin and aliquot. Reconstitution concentration should be appropriate to dilute to 100 ng/mL in cell media. Store aliquots at 20
C for 3 months (see Note 3). 22. BMP4: Reconstitute in 4 mM HCl supplemented with 0.1% bovine serum albumin and aliquot. Reconstitution concentration should be appropriate to dilute to 10 ng/mL in cell media. Store aliquots in 80
C for 3 months (see Note 3). 23. VEGF165: Reconstitute in deionized water and aliquot. Reconstitution concentration should be appropriate to dilute to 50 ng/mL in media. Store aliquots at 20
C for 3 months (see Note 3). 24. RPMI 1640 + B27 medium: RPMI1640 medium with 2 mM L-glutamine and 2% B27 supplement without insulin. 25. RPMI 1640 + FBS medium: RPMI 1640 medium with 10% FBS and 2 mM L-glutamine. 26. MEF medium composed of DMEM with 10% fetal calf serum, 2 mM L-glutamine, 1 nonessential amino acids, penicillinstreptomycin 10,000 U/mL (ThermoFisher Scientific, Gibco™, catalog number: 15140-122,). Store at 4
C. 27. Embryonic stem cell medium: knockout DMEM with 20% knockout serum replacement, 1 mM L-glutamine, 0.1 mM 2-mercaptoethanol, 1 nonessential amino acids, 4 ng/mL hbFGF. Store at 4
C. 28. MEF conditioned medium (MEF-CM): Treat MEF cells with 10 μg/mL mitomycin C for 2.5 h, harvest, and seed at a density of 55,000 cells/cm2 in MEF medium. After 1 day,

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exchange MEF medium for embryonic stem cell medium [18]. Collect media daily for 1 week [18]. Store aliquoted MEF-CM at 20
C after filtration; use within 1 month (see Notes 4 and 5). Add 4 ng/mL hbFGF to the MEF-CM immediately before adding to the cell plate [18]. 29. CTK solution: 0.1% collagenase IV, 0.25% Trypsin, 20% knockout serum replacement (KSR), and 1 mmol/L CaCl2 in phosphate-buffered saline (PBS). 30. CO2 incubator: 37
C, 5% CO2, humidity pan filled with distilled water. 2.2 Antibodies and Staining for Cell Lineage Analysis

1. Antibodies used for cell lineage analysis: anti-PDGFRβ conjugated with phycoerythrin, anti-VE cadherin conjugated with fluorescein isothiocyanate (FITC), anti-cardiac isoform of troponin T (cTnT) conjugated with Alexa-488, and anti-TRA-160 conjugated with FITC. Store all antibodies undiluted at 20
C. Antibody expiration is lot specific (see Note 6). 2. LIVE/DEAD Aqua dead cell staining kit: Store kit at 20
C. Stable for 6 months (see Note 4). 3. PBS without Ca2+ and Mg2+. 4. 16% paraformaldehyde, methanol free. 5. Saponin. 6. 4% paraformaldehyde in PBS. 7. FACS staining buffer: PBS with 5% FBS. 8. FACS staining buffer for anti-cTnT: PBS with 5% FBS and 0.75% saponin. 9. BD LSRII Flow Cytometer with DIVA software or equivalent replacement.

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ECT Construction

1. High glucose DMEM: Store at 4
C. 2. FBS: Store at 20
C. 3. Alpha minimum essential medium (αMEM; purchased, show components?): Store at 4
C. 4. 100 penicillin-streptomycin: Store at 20
C. 5. Tissue culture grade 2-mercaptoethanol: Store at 4
C. 6. Accumax: Store at 20
C. Use within 2 months after thawing. 7. Acidic rat tail collagen type I: Store at 4
C for 12 months. 8. Growth factor-reduced Matrigel: Store 100 μL aliquots at 20
C. 9. High glucose DMEM with 20% FBS: Store at 4
C.

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10. ECT medium: αMEM with 10% FBS, 1 penicillinstreptomycin, and 5  105 M 2-mercaptoethanol. Store at 4
C. 11. Alkali buffer: distilled water with 200 mM NaHCO3, 200 mM HEPES, and 100 mM NaOH. Store at 4
C. Replace buffer every month. 12. (AAV)1/2-CAG-oChIEF-tdTomato vector or similar optogenetic viral vector (see Note 7). 13. CO2 incubator: 37
C, 5% CO2, humidity pan filled with distilled water. 14. Flexcell® FX-5000™ Tissue Train® system and collagen type Iand IV-coated silicone membrane 6-well Linear Tissue Train® plates: Install the Tissue Train® system in a CO2 incubator (see Note 8). 2.4 Chronic Light Stimulation

1. Luxeon Rebel 470 nm LEDs (see Note 9). 2. Fraen OMG 9
22 mm circular beam lens: one per LED. 3. Heat sink: must be large enough to hold the Tissue Train® 6-well plate. 4. BuckPuck 3021-D-I-1000 LED driver: one per LED. 5. Thermal tape. 6. Silicone adhesive. 7. 22-gauge hook-up wire. 8. Wire stripper and cutter. 9. 7-pin MTA-100 connector for 22-gauge wire: one connector per LED. 10. Strain relief cover for 7-pin MTA-100: one cover per LED. 11. Crimping or insertion tool for MTA-100 connector. 12. Male and female barrel connectors: three connectors per LED. 13. Soldering iron. 14. Solder. 15. Heat shrink tubing. 16. Power supply: The LED driver operates between 7 and 32 V. Each LED driver draws 1 A of current. Depending on the number of LEDs you plan to connect, make sure your power supply can support the current and voltage requirements. 17. Arduino Uno, supplied with USB cable. 18. DC wall adapter for Arduino USB cable. 19. Arduino IDE software (free download from Arduino website). 20. Arduino sketch to pace ECT constructs (Subheading 3.8).

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21. 37
C incubator with a video microscope capable of focusing on the ECT. We used an Mu1000 AmScope with ToupView software. 2.5 Mechanical Force Testing

1. 2,3-Butanedione monoxime (BDM). 2. Tyrode’s solution: distilled water with 120 mM NaCl, 5.4 mM KCl, 2.5 mM CaCl2, 1 mM MgCl2, 22.6 mM NaHCO3, 0.4 mM NaH2PO4, 0.05 mM Na2EDTA, 0.28 mM ascorbic acid, and 0.56 mM glucose. Bubble Tyrode’s solution with 95/5% O2/CO2 gas. 3. Tyrode’s solution with 30 mM BDM. 4. 10-0 nylon suture. 5. Microdissection scissors. 6. Microdissection forceps (#55). 7. Force testing apparatus: brightfield microscope, temperatureregulated perfusion chamber, force transducer (model 801C, Aurora Scientific), micromanipulator, and pacing electrodes. A desktop computer with suitable software is needed to display readings from the force transducer.

3

Methods Carry out procedures at room temperature unless otherwise specified.

3.1 Differentiation and Maintenance of hIPSCs

1. Coat cell culture plate with thin-coat Matrigel. Use enough to cover the bottom of the dish. Leave coated plate for 1 h at room temperature. 2. Plate h-iPSCs at a plating density appropriate for the cell culture dish being used. Add the appropriate amount of MEF-CM with 1 ng/mL hbFGF for your cell culture dish, and expand the cells. 3. Passage cells every 4–6 days using CTK solution. Remove MEF-CM. Wash cells with Ca2+ and Mg2+ free PBS. Add enough CTK solution to cover the bottom of the cell culture dish. Incubate at 37
C for 5 min. Remove the dish and transfer contents to a 15 mL conical tube. Spin down cells at an appropriate speed for your cell type. Remove the CTK solution and add fresh MEF-CM with 4 ng/mL hbFGF to cells. Break up the cells with 1000 mL pipette tip into small clusters, and passage clusters into new culture dishes.

3.2 Cardiovascular Differentiation

1. Use Versene to detach and passage cells when confluency is reached. Plate cells at a density of 10,000 mm2 on thin-coat

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Matrigel-coated cell culture dishes in MEF-CM with 4 ng/mL hbFGF, and culture for 2–3 days. 2. Coat cells with thin-coat Matrigel for 1 day. 3. Replace the MEF-CM with RPMI 1640 + B27 medium. Add 100 ng/mL of Activin A and 100 ng/mL of Wnt3A to the medium for 1 day. This is day 0 of differentiation. 4. On day 1 of differentiation, change to new RPMI 1640 + B27 medium with 10 ng/mL of BMP4 and 10 ng/mL of hbFGF. Culture cells for 2 or 4 days with no media change. 5. To induce cardiomyocyte and endothelial differentiation (CM + EC protocol), change the RPMI 1640 + B27 medium on day 5 of differentiation, and supplement with 50 ng/mL VEGF165. Change the culture medium every 48 h (see Note 10). 6. To induce mural cell differentiation, replace the medium with RPMI1640 + FBS at day 3 of differentiation, and change the medium every 48 h (see Note 10). 3.3

Lineage Analysis

1. On differentiation day 15, harvest the cells from cell culture plates using Accumax. Wash the cells with Ca2+ and Mg2+ free PBS. Pipette enough Accumax in the cell culture dish to cover the plate. Incubate the plate for 5 min at 37
C. Harvest the cells after incubation (see Note 11). 2. To eliminate dead cells, cells were stained with the LIVE/ DEAD fixable Aqua dead cell staining kit (Life Technologies). 3. For cell surface markers, staining was carried out in PBS with 5% FBS. 4. For intracellular proteins, staining was carried out on cells fixed with 4% paraformaldehyde (PFA) in PBS. 5. Resuspend and fix 1  106 of the day 15 harvested cells in 4% paraformaldehyde and stain with anti-cTnT. Dilute 1:50 in FACS staining buffer for anti-cTnT. After antibody staining, stain with the LIVE/DEAD Aqua dead cell staining kit (Life Technologies) according to manual instructions. Harvest the stained cells into FACS tubes, and analyze with an LSRII flow cytometer with DIVA software. 6. Harvest 1  106 cells and stain with membrane surface markers. Use the following dilutions of antibody in FACS staining buffer: anti-PDGFRβ (1:100), anti-VE cadherin (1:100), and anti-TRA-1-60 (1:20). Analyze the stained cells with an LSRII flow cytometer with DIVA software (see Note 12).

3.4 Construction of ECTs

1. For each ECT, resuspend 3  106 of the harvested cells from the CM + EC and MC protocols in 83 μL of high glucose DMEM with 20% FBS in a sterile 15 mL Eppendorf tube (see

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Note 13). Use a cell mixture of approximately 60% CM, 20% EC, and 20% MC. 2. On ice in a separate PCR tube, neutralize 33 μL of acidic rat tail collagen with 54 μL of alkali buffer. 3. Finally, add 30 μL of Matrigel to the collagen tube. Add the collagen/Matrigel to the cell population to make a total solution volume of ~200 μL (see Note 14). 4. Add the (AAV)1/2-CAG-oChIEF-tdTomato vector to the cell mixture at a MOI of 500 (see Note 7). 5. Plate the cell mixture into the Tissue Train® plate. Push the anchor tabs down on either side of the well, and plate the tissue mixture such that both ends are touching the tabs on opposite sides of the well. Load the plate into the Tissue Train® system,
and apply vacuum suction for 2 h in the incubator at 37 C, 5% CO2 (see Note 14). 6. After 2 h, turn off the vacuum suction, remove the plate from the Tissue Train® system, and add 2 mL of ECT medium.
Continue to culture at 37 C, 5% CO2. Change the medium every other day (see Note 15). 3.5 Construction of LED Array

1. Program the Arduino Uno using the sketch supplied in Subheading 3.8 (see Notes 16 and 17). 2. Use thermal tape to mount LEDs onto the heat sink. Mount LEDs such that they are centered at the positions of the wells of the Tissue Train® 6-well plate. 3. Solder wire to the terminals of a female barrel connector. Solder the other ends to the terminals of the LED. Repeat this for each LED on your heat sink. 4. Attach a circular beam lens to each LED with silicone adhesive. 5. For each LED, solder wires to the terminals of a female barrel connector, and connect the other ends to your power supply (see Note 18). Connect multiple LEDs in parallel to the power supply. 6. For each LED, solder wires to the terminals of a female barrel connector. Connect one wire to one of the pulse wave modulation (PWM) digital I/O pins of the Arduino Uno (see Note 19). Connect the other wire to a ground pin of the Arduino Uno. The Arduino Uno has three ground pins. Connect up to two LEDs per pin. Use solder and heat shrink tubing to join two wires together. 7. For each LED, make the wire connections to an LED driver shown in Fig. 1. Attach an MT-100 connector to the LED driver, and use the crimping or insertion tool to connect the wiring as shown.

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Fig. 1 LED wiring diagram, assembled LED circuit with a single LED, and force testing apparatus. (a) Single LED wiring diagram. Connect to LED driver using an MTA-100 connector. Wires are soldered to barrel connector terminals. Connect the LED () terminal to the Arduino GND pin using heat shrink tubing. (b) Representative LED assembly, showing the connections between the Arduino microcontroller, LED driver, and LED. (c) Force testing apparatus with force transducer, pacing electrode, perfusion injection ports, length controller, and micromanipulator indicated

8. Connect the barrel connector between the LEDs and the LED drivers. 9. Connect the barrel connectors between the Arduino and the LED drivers. Be sure you know which Arduino I/O pin controls which LED. 10. Plug in the Arduino so that the program is running. 11. Connect the barrel connectors between the LED drivers and the power supply. If the power supply is not plugged in or turned on, do so now. The LEDs will begin to pulse (see Note 20). 12. The Tissue Train® plate cannot be placed directly on top of the LEDs. You will need an approximately 25-mm-tall insert placed on top of the heat sink, upon which you can place the Tissue Train® plate containing the ECTs. We used a Tissue Train® plate for this purpose. We removed the bottom portion and the silicone membrane and placed the plate upside-down onto our heat sink (Fig. 1b). We painted the plate matte black to optically isolate the LEDs. Additionally, we cut small notches for the wires using a Dremel tool.

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3.6 Chronic Light Stimulation

1. Seven days after ECT formation, begin C-OP of transfected tissue (see Note 20). 2. Place the ECT plate over the LED array/heat sink in a 37
C incubator containing a video microscope. Run the wires out of the incubator. Place the Arduino Uno, LED driver, and power supply in a location such that the wires are not tugging on the LED components in the incubator. Be sure you have placed the components such that you can plug the Arduino Uno into a laptop and the power supply into a wall outlet. 3. Plug the Arduino Uno into a laptop with the Arduino IDE installed, and plug the power supply into a nearby wall outlet (see Note 21). 4. Open the Arduino software and sketch on the laptop. 5. Position an ECT underneath the microscope. 6. Focus on the ECT with the microscope and switch to the video feed. 7. Record the intrinsic beat rate of the ECT. Manually count, through observation of the video feed, the ECT beat rate to obtain the intrinsic frequency. 8. In the Arduino sketch, change the pacing frequency of the appropriate I/O pin to 0.5 Hz above the intrinsic beat rate. Upload the new sketch into the Arduino microcontroller. Manually count to ensure optical capture by the ECT. 9. Continue to increase the optical pacing rate by 0.5 Hz until the ECT can no longer optically capture at the set pacing frequency. This frequency is considered the optical maximum capture rate (O-MCR) for that day. 10. Change the Arduino sketch for the appropriate I/O pin such that the pacing frequency is 0.5 Hz below the O-MCR. This frequency will be used for chronic pacing. Upload the new sketch into the Arduino Uno microcontroller. 11. Repeat steps 5–10 for each ECT. 12. Sterilize the LED pacing system and tissue plate by gently wiping all components that will be placed in the incubator with 70% ethanol. 13. Position the ECT plate and pacing system in your long-term incubator, in a similar fashion as described for the short-term optical capture above. The only difference is to make sure you are close enough to an outlet to plug in the Arduino with a DC wall adapter (see Note 21). For additional light isolation, cover the lid of the ECT plate with aluminum foil during chronic pacing.

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14. The next day, and through day 13 after ECT generation, repeat O-MCR recording, and reset the pacing frequency every day to 0.5 Hz below the O-MCR. 3.7

Force Testing

1. On day 14 after ECT formation, fill the perfusion chamber of the force testing system with Tyrode’s solution supplemented with BDM, and preheat the solution in the perfusion chamber to 37
C. 2. Prebubble ~250 mL of Tyrode’s solution with 95/5% O2/ CO2 and warm to 37
C. 3. Cut the ECT from the Tissue Train® plate using a small pair of sterile surgical scissors. Cut the ECT, making sure not to leave the anchor tabs attached to the tissue. Place the ECT, using small sterile forceps, into a 35 mm dish filled with the warmed Tyrode’s supplemented with BDM. 4. Transfer the ECT into the perfusion chamber. 5. Using small sterile forceps and 10-0 suture, tie the tissue to the force transducer and length controller in the perfusion chamber (see Note 22). 6. Start perfusing with warmed Tyrode’s solution, prebubbled in step 2. Perfuse with Tyrode’s solution for 20 min to wash out the BDM. 7. Open the software on the computer for force recordings. Begin recording active force. 8. Use the micromanipulator attached to the length controller to stretch the ECT by 0.5 mM increments. 9. Keep stretching the ECT until active force does not increase. This is the Lmax of the tissue. Perform all further force recordings at Lmax. 10. Take an image at Lmax to calculate ECT diameter during analysis. 11. Perfuse more Tyrode’s solution into the chamber for 5 min. 12. Record the intrinsic beat rate of the ECT. 13. Connect an unmounted LED to the Arduino circuit, and place the LED underneath the perfusion chamber. 14. Use the Arduino microcontroller to optically pace the tissue 0.5 Hz above the intrinsic beat rate. Keep increasing the optical pacing rate by 0.5 Hz until the O-MCR is reached (Fig. 2). 15. Perfuse Tyrode’s solution into the chamber for 5 min and remove the LED circuit. 16. Begin force recording again and record the intrinsic beat rate of the ECT. 17. Begin electrical pacing at 2 Hz for 20 min.

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18. Using electrical pacing, increase the pacing rate in 0.5 Hz increments until the MCR is reached (Fig. 3). 19. Save images and force recordings for analysis. 20. Using an image processing software and the ECT image at Lmax, measure the ECT diameter in three different locations along the tissue. Average these measurements to obtain the mean diameter. Calculate the cross-sectional area, using an equation for circular area. 21. Measure the following parameters for each recording and, when applicable, every pacing frequency: intrinsic beat rate, O-MCR, electrical pacing MCR, mean active stress (σ A), relaxation time to 50% diastolic stress (RT50), beat-to-beat hysteresis (σ bb), and systolic potential energy (Fig. 4). 3.8

Arduino Sketch

// Pulse an LED at a given frequency and pulse duration // Edit these parameters to control the LED pulses

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Fig. 3 Recording and analysis of MCR during electrical pacing of a transfected tissue. (a) Recording showing MCR capture at 5.5 Hz during electrical ramp-up pacing protocol. (b) Fast Fourier transform (FFT) analysis for verification that tissue captured at 5.5 Hz electrical pacing frequency. (c) Capture at 6 Hz electrical pacing does not occur. (d) FFT analysis verifying that electrical capture does not occur at 5 Hz // Only edit the number after the equals sign. Nothing else. const double freq3 ¼ 2.5; //frequency of pin 3 in Hz const double p3 ¼ 5; //pulse duration of pin 3 in ms const double freq5 ¼ 2.5; //frequency of pin 5 in Hz const double p5 ¼ 5; //pulse duration of pin 5 in ms const double freq6 ¼ 2.5; //frequency of pin 6 in Hz const double p6 ¼ 5; //pulse duration of pin 6 in ms const double freq9 ¼ 3.5; //frequency of pin 9 in Hz const double p9 ¼ 5; //pulse duration of pin 9 in ms const double freq10 ¼ 3.75; //frequency of pin 10 in Hz const double p10 ¼ 5; //pulse duration of pin 10 in ms const double freq11 ¼ 3.5; //frequency of pin 11 in Hz const double p11 ¼ 5; //pulse duration of pin 11 in ms // Edit these constants if the PMW pins on your Arduino do not match the ones in this sketch // for example, if pin 3 on your board is not a PMW pin, but pin 4 is, you can change // “const int ledPin3 ¼ 3;” to “const int ledPin3 ¼ 4;”

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Fig. 4 Typical analysis of a stress-time recording. Active stress is the mean average distance between systolic and diastolic force, normalized to ECT cross-sectional area. Red dots mark peak systolic stress and blue dots mark end diastolic stress. Beat-to-beat variation is the average difference between the active stress of two successive beat cycles. RT50 is the relaxation time to 50% of the maximum force for a cycle. PE is the summation of the systolic stress-time integral // now all the settings for pin 3 above will actually run through pin 4 // Only edit the number after the equals sign. Nothing else. const int ledPin3 ¼ 3; const int ledPin5 ¼ 5; const int ledPin6 ¼ 6; const int ledPin9 ¼ 9; const int ledPin10 ¼ 10; const int ledPin11 ¼ 11; // do not edit anything below this line unsigned long period3 ¼ 1000/freq3; unsigned long tf3 ¼ 0; int ledState3 ¼ LOW; unsigned long period5 ¼ 1000/freq5; unsigned long tf5 ¼ 0; int ledState5 ¼ LOW; unsigned long period6 ¼ 1000/freq6; unsigned long tf6 ¼ 0;

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int ledState6 ¼ LOW; unsigned long period9 ¼ 1000/freq9; unsigned long tf9 ¼ 0; int ledState9 ¼ LOW; unsigned long period10 ¼ 1000/freq10; unsigned long tf10 ¼ 0; int ledState10 ¼ LOW; unsigned long period11 ¼ 1000/freq11; unsigned long tf11 ¼ 0; int ledState11 ¼ LOW; void setup() { pinMode(ledPin3,OUTPUT); pinMode(ledPin5,OUTPUT); pinMode(ledPin6,OUTPUT); pinMode(ledPin9,OUTPUT); pinMode(ledPin10,OUTPUT); pinMode(ledPin11,OUTPUT); } void loop() { unsigned long t ¼ millis(); //current time since program began tf3 ¼ t%period3; //cycle time for pin 3 in ms if (tf3