Stem Cell Renewal and Cell-Cell Communication: Methods and Protocols (Methods in Molecular Biology, 2346) [2nd ed. 2021] 1071615696, 9781071615690

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Stem Cell Renewal and Cell-Cell Communication: Methods and Protocols (Methods in Molecular Biology, 2346) [2nd ed. 2021]
 1071615696, 9781071615690

Table of contents :
Preface
Contents
Contributors
Inference of Ligand-Receptor Pairs from Single-Cell Transcriptomics Data
1 Introduction
2 Materials
2.1 Input Data Files
2.2 Software
2.3 Hardware
3 Methods
3.1 Installation
3.2 Running in the Statistical Analysis Mode
3.3 Subsampling and Statistical Analysis
3.4 Normal Mode, Without Statistical Analysis
3.5 Visualization
3.6 Different Versions of the CellPhoneDB Database
3.7 Generating a User-Specific Database
3.8 Help Option
3.9 Interactive Web Portal
4 Notes
References
Multiple Imaging Modalities for Cell-Cell Communication via Calcium Mobilizations in Corneal Epithelial Cells
1 Introduction
2 Materials
3 Methods
3.1 Preparation of Cells for Ca2+ Mobilization Studies
3.2 Ex Vivo In Situ Preparation of Tissue for the Study of Calcium Mobilizations
3.3 Imaging
3.4 Analysis
3.4.1 Grid-Based Modeling of Ca2+ Mobilizations
3.4.2 Cell-Based Modeling of Ca2+ Mobilizations
3.4.3 Event Probability Calculation for Ca2+ Mobilizations
3.4.4 Analysis of Cytoskeletal Changes with Calcium Mobilizations
4 Notes
References
Interactions of Hematopoietic Stem Cells with Bone Marrow Niche
1 Introduction
2 Materials
2.1 Tissues
2.2 Reagents and Supplies
2.3 Fluorescence-Conjugated Antibodies
2.4 Equipment
3 Methods
3.1 Preparing Single-Cell Suspension from Long Bones (Femur and Tibia)
3.2 Staining BM Cells and Endosteal Stromal Cells for Flow Cytometry Analysis
3.3 Staining the Endosteal Stromal Cells for Flow Cytometry Sorting
3.4 HSCs and Stromal Cells Co-culture
4 Notes
References
Ex Vivo Modeling of Hematopoietic Stem Cell Homing to the Fetal Liver
1 Introduction
2 Materials
2.1 Microfluidic Channel
2.2 Polyacrylamide Hydrogels
2.3 Collagen Gel Coating
2.4 Fetal Liver Tissue
2.5 AGM Dissociation for Isolation of Hematopoietic Stem and Progenitor Cells
2.6 Fluidics Assembly and Imaging
3 Methods
3.1 Microfluidic Device Assembly
3.1.1 Microfluidic Channel Base Slide
3.1.2 Polyacrylamide Hydrogels
3.1.3 Collagen Coating
3.2 Fetal Liver-on-a-Chip Assembly
3.3 HSC Isolation
3.4 Fluidics Assembly and Imaging
4 Notes
References
Analysis of Epithelial Architecture and Planar Spindle Orientation in the Drosophila Wing Disc
1 Introduction
2 Materials
2.1 Dissection of Wing Discs
2.2 Immunostaining of Wing Discs
2.3 Mounting Wing Discs on Slides
2.4 Imaging Wing Discs
2.5 Angle Analysis of Mitotic Spindle Orientation
3 Methods
3.1 Dissection of Larvae
3.2 Immunostaining
3.3 Mounting of Wing Imaginal Discs
3.4 Image Acquisition
3.5 Visualization of Epithelial Architecture
3.6 Measurements of Planar Spindle Orientation
4 Notes
References
A Co-culture Model to Study the Effect of Kidney Fibroblast-p90RSK on Epithelial Cell Survival
1 Introduction
2 Materials
2.1 Mice
2.2 Cell and Cell Culture Reagents
2.3 Other Instruments and Materials
3 Methods
3.1 Kidney Fibroblast Isolation
3.2 Culture of Fibroblasts
3.3 Culture of Epithelial Cells
3.4 Co-culture of Epithelial Cells and Fibroblasts
4 Notes
References
Calcium Fluorescence Recordings from Neuroepithelial Stem Cells
1 Introduction
2 Materials
2.1 Recording Apparatus
2.2 Fluorescent Ca2+ Indicators
2.3 Solutions
3 Methods
3.1 Preparation of Retinal Neuroepithelium
3.2 Loading of Ca2+ Indicator
3.3 Ca2+ Fluorescence Measurement
4 Notes
References
Ultrastructural Analysis of Cell-Cell Interactions in Drosophila Ovary
1 Introduction
2 Materials
2.1 Sample Preparation
2.2 ROTO Processing
2.3 Resin Preparations and Embedding
2.4 Sectioning
3 Methods
3.1 Fly Culture
3.2 Sample Preparation
3.3 Processing
3.3.1 First Osmium Penetration
3.3.2 TCH Incubation
3.3.3 Second Osmium Infiltration
3.3.4 Lead Aspartate Staining
3.3.5 Dehydration
3.3.6 Resin Infiltration
3.4 Embedding
3.5 Sectioning
3.5.1 Trimming the Block
3.5.2 Ultramicrotome
3.5.3 Section Preparation and Collection
3.5.4 Imaging
4 Notes
References
TIRF Microscopy as a Tool to Determine Exosome Composition
1 Introduction
2 Materials
2.1 Cells
2.2 Reagents
2.3 Media and Solutions/Buffers
2.4 Equipment
2.5 Software
3 Methods
3.1 CD4+ T-Cell Purification from Human Peripheral Blood Lymphocytes (huPBLs)
3.2 Generation of Lymphoblasts from Human PBLs: PHA or SEE-Specific
3.3 Isolation of EVs from Cell Culture
3.3.1 Production and Isolation of EVs (Fig. 1a)
3.3.2 Quantification and Characterization of EVs
3.4 Coating of Coverslips and Coverslip-Bottom Dishes
3.4.1 Preparation of Poly-L-Lysine-Coated Surfaces
3.4.2 Preparation of Antibody-Coated Surfaces
3.5 Sample Preparation for Imaging
3.5.1 Preparation of Samples on Poly-L-Lysine-Coated Surfaces (Fig. 1b)
3.5.2 Preparation of Samples on Capture Antibodies-Coated Surfaces(Fig. 1c)
3.6 Imaging and Analysis Under TIRF Microscope
4 Notes
References
Rapid Visualization of Intracellular Vesicle Events During Synaptic Stimulation
1 Introduction
2 Materials
2.1 Primary Cells and Cell Lines
2.2 Reagents
2.3 Media
2.4 Equipment
2.5 Software
3 Methods
3.1 Isolation of T Cells
3.1.1 Isolation of Human PBLs
3.1.2 Isolation of Mouse CD4+ T Cells
3.2 Generation of SEE-Specific or PHA Lymphoblasts from Human PBLs
3.3 Transfection of T Cells
3.3.1 Electroporation of Lymphoblastoid Cell Lines and Human SEE T Lymphoblasts
3.3.2 Nucleofection of Human T Lymphoblasts
3.3.3 Nucleofection of Mouse CD4+ T Cells
3.4 Coating of Coverslip-Bottom Chambers
3.4.1 Preparation of poly-L-Lys-Coated Surfaces
3.4.2 Preparation of Stimulating Surfaces
3.5 Live Imaging Vesicular Traffic Studies
3.5.1 Preparation of T Cells
3.5.2 Image Acquisition
3.5.3 Image Analysis LAS X Software (See Fig. 1)
3.5.4 Image Analysis Imaris Bitplane Software (See Fig. 2)
4 Notes
References
Monitoring of Active Notch Signaling in Mouse Bladder Urothelium
1 Introduction
2 Materials
2.1 Mice
2.2 Tissue Fixation Reagents and Components
2.3 Paraffin Embedding Reagents and Equipment
2.4 Immunostaining Reagents
2.5 Other Equipment for Immunostaining
2.6 Microscopy and Imaging Set Up
2.7 Image Processing and Analysis
3 Methods
3.1 Tissue fixation
3.2 Paraffin Embedding and Sectioning Onto Slides
3.3 Immunostaining on Paraffin Sections for Detection of Nuclear Notch ICD Levels
3.3.1 Deparaffinization and Rehydration of Tissue
3.3.2 Antigen Retrieval
3.3.3 Blocking
3.3.4 Primary Antibody Binding
3.3.5 Secondary Antibody Binding
3.3.6 Tyramide Signal Amplification
Cover-Slipping
Imaging
4 Notes
References
Examining Local Cell-to-Cell Signalling in the Kidney Using ATP Biosensing
1 Introduction
2 Materials
2.1 Tissue Culture
2.2 Instrumentation
2.3 Balanced Salt Solution (BSS)
2.4 Biosensors
3 Methods
3.1 Tissue Culture
3.2 Initial Rig Preparation
3.3 Experiment Preparation
3.4 Biosensor Polarization
3.5 Experiment
3.6 Analysis
4 Notes
References
Isolation and Assessment of Pancreatic Islets Versus Dispersed Beta Cells: A Straightforward Approach to Examine Cell-Ce
1 Introduction
2 Materials
2.1 Islet Isolation
2.2 Islet Dispersion
2.3 Calcium Imaging
2.4 Islet Cell Staining
3 Methods
3.1 Islet Isolation
3.2 Islet Dispersion
3.3 Calcium Imaging
3.4 Islet Cell Staining
4 Notes
References
Promoter Pull-Down Assay: A Biochemical Screen for DNA-Binding Proteins
1 Introduction
2 Materials
2.1 Probe Preparation
2.2 Buffers and Reagents
2.3 Cell Lysis
2.4 Pull-Down
2.5 SDS-PAGE Gel
3 Methods
3.1 Cell Culture
3.2 Probe Preparation
3.3 Dynabead Preparation
3.4 Probe Immobilization
3.5 Bead Equilibration
3.6 Cell Lysis
3.7 Preclearing the Lysate
3.8 Dynabead Incubation with Lysate
3.9 Elution
3.10 Analysis of Eluted Proteins
4 Notes
References
Purification of the Vibrio Quorum-Sensing Transcription Factors LuxR, HapR, and SmcR
1 Introduction
2 Materials
2.1 Cell Culture Reagents
2.2 Buffers and Reagents
2.3 Cell Lysis
2.4 Protein Purification
2.5 SDS-PAGE Gel
2.6 Autoinduction Media (Optional)
3 Methods
3.1 Protein Overexpression (See Note 2)
3.2 Cell Lysis
3.3 Protein Purification: Native Purification (See Note 7)
3.4 Protein Purification: Affinity Tag Option 1 - His-Tag (See Note 10)
3.5 Protein Purification: Affinity Tag Option 2 - IMPACT (See Note 11)
3.6 Protein Purification: Size Exclusion Chromatography (SEC) (See Note 13)
4 Notes
References
Preserving Cytonemes for Immunocytochemistry of Cultured Adherent Cells
1 Introduction
2 Materials
2.1 Cell Culture
2.2 Fixation, Permeabilization and Block
2.3 Immunocytochemistry
2.4 Mounting Coverslips
3 Methods
3.1 Cell Culture
3.2 Fixation, Permeabilization and Blocking
3.3 Antibody Binding
3.4 Mounting
4 Notes
References
Fluorescent Labeling of Connexin with As Complex and X-Y Coordinate Registration of Target Single Cells Based on a Triangle St
1 Introduction
2 Materials
2.1 HeLa Cell Culture
2.2 Transfection
2.3 Fluorescent Dyes for Diffusion Marker
2.4 Chelating Reagent for a TC Tag and Washing Buffer
2.5 Polystyrene Plate
2.6 Cell Fixation Reagents
3 Methods
3.1 Preparation of an Insert DNA: A Cx43-TC Chain
3.2 Isolation of a Plasmid Vector pDsRed Monomer C1
3.3 Linearization of the Plasmid Vector
3.4 Ligation of Cx43-TC Chain and the Vector to Generate pCMV-Cx43-TC
3.5 Transfection of HeLa Cells and FlAsH Staining
3.6 Analysis of GJ Plaque Localization and Gap Junctional Dye Diffusion
3.7 Preparation of a Culture Dish with a Coordinate Chip
3.8 Correlation of X-Y Coordinates Determined by Light and Electron Microscopy
4 Notes
References
Chemical and Voltage Gating of Gap Junction Channels Expressed in Xenopus Oocytes
1 Introduction
2 Materials
2.1 Solutions Used During Oocyte Preparation and in Electrophysiological Experiments
3 Methods
3.1 Oocyte Preparation
3.2 Inhibition of Native-Connexin Expression
3.3 Inhibition of Calmodulin (CaM) Expression
3.4 Expression of CaM Mutant
3.5 Oocyte Injection of Connexin cRNAs and Oocyte Pairing
3.6 Measurement of Gap Junctional Conductance by Double Voltage Clamp
3.7 Measurement of Trans Junctional Voltage Gating
3.8 Oocyte Chamber Perfusion and Uncoupling Protocol
3.9 Bubble-Trap Device
4 Notes
References
The Analysis of Gap Junctional Intercellular Communication Among Osteocytes in Chick Calvariae by Fluorescence Recovery After
1 Introduction
1.1 Osteocyte Network
1.2 Fluorescence Recovery After Photobleaching
2 Materials
2.1 Animals
2.2 General Reagents
2.3 Equipment
3 Methods
3.1 Tissue Collection and Calcein AM Loading
3.2 FRAP Experiment on an Osteocyte
3.3 Data Analyses
4 Notes
References
Flow Cytometry Evaluation of Gap Junction-Mediated Intercellular Communication Between Cytotoxic T Cells and Target Tumor Cell
1 Introduction
2 Materials
2.1 Common Reactive and Equipment
2.2 Melanoma-Associated Antigen (gp10025-33)-Specific CTL Generation
2.3 Wild-Type Naive CD8+ T Cell Generation
2.4 Calcein-Acceptor Tumor Cells
2.5 Calcein-Donor Cell Labeling
2.6 Calcein-Acceptor Cell Labeling
2.7 Co-incubation of Calcein-Donor and Calcein-Acceptor Cells
2.8 Flow Cytometry
3 Methods
3.1 Melanoma Antigen gp100-Specific CTL Generation
3.2 Wild-Type Naive CD8+ T Cell Obtaining
3.3 Calcein-Acceptor Tumor Cells Obtaining
3.4 Calcein-Donor Cell Labeling
3.5 Calcein-Acceptor Cell Labeling
3.6 Co-incubation of Calcein-Donor and Calcein-Acceptor Cells
3.7 Flow Cytometry Analysis
4 Notes
References
Index

Citation preview

Methods in Molecular Biology 2346

Kursad Turksen Editor

Stem Cell Renewal and Cell-Cell Communication Methods and Protocols Second Edition

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 Pub Med.

Stem Cell Renewal and Cell-Cell Communication Methods and Protocols Second Edition

Edited by

Kursad Turksen Ottawa, ON, Canada

Editor Kursad Turksen Ottawa, ON, Canada

ISSN 1064-3745 ISSN 1940-6029 (electronic) Methods in Molecular Biology ISBN 978-1-0716-1569-0 ISBN 978-1-0716-1570-6 (eBook) https://doi.org/10.1007/978-1-0716-1570-6 © 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. Cover Illustration Caption: Artwork by Kursad Turksen. 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 Great strides have been made in the field of cell–cell communications with respect to the identification and characterization of key components of the communication apparatus, assembly and maintenance of the communications structures, and concomitantly their roles in not only tissue formation and maintenance but also regeneration and repair. In this second edition of the Stem Cell Renewal and Cell-Cell Communication volume, I have brought together a new set of protocols to arm cell biologists with protocols that are currently being used in a number of well-established laboratories around the world. I hope that both people already in the field as well as newcomers will benefit from this compilation, and that the volume will drive continued growth in our understanding of the crucial biological and physiological roles of cell–cell communications in tissue function and organismal integrity. Once again, the protocols gathered here are faithful to the mission statement of the Methods in Molecular Biology series: They are well-established and described in an easy to follow, step-by-step fashion so as to be valuable for not only experts but also novices in the field. That goal is achieved because of the generosity of the contributors who have carefully described their protocols in this volume, and I am very grateful for their efforts. My thanks as well go to Dr. John Walker, the Editor-in-Chief of the Methods in Molecular Biology series, for giving me the opportunity to create this volume and for supporting me along the way. I am also grateful to Patrick Marton, the Executive Editor of Methods in Molecular Biology and the Springer Protocols collection, for his continuous support from idea to completion of this volume. A special thank you goes to Anna Rakovsky, Assistant Editor for Methods in Molecular Biology, for her continuous support from beginning to end of this project. I would also like to thank David C. Casey, Senior Editor of Methods in Molecular Biology, for his outstanding editorial work during the production of this volume. Finally, I would like to thank Anand Ventakachalam and the rest of the production crew for their work in putting together an outstanding volume. Ottawa, ON, Canada

Kursad Turksen

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

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Inference of Ligand–Receptor Pairs from Single-Cell Transcriptomics Data. . . . . . . . . Mirjana Efremova and Roser Vento-Tormo Multiple Imaging Modalities for Cell-Cell Communication via Calcium Mobilizations in Corneal Epithelial Cells . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Yoonjoo K. Lee, Kristen L. Segars, and Vickery Trinkaus-Randall Interactions of Hematopoietic Stem Cells with Bone Marrow Niche. . . . . . . . . . . . . . . Xinghui Zhao, Cuiping Zhang, Xiaojing Cui, and Ying Liang Ex Vivo Modeling of Hematopoietic Stem Cell Homing to the Fetal Liver . . . . . . . . . Amina Mohammadalipour, Miguel F. Diaz, Sumedha Pareek, and Pamela L. Wenzel Analysis of Epithelial Architecture and Planar Spindle Orientation in the Drosophila Wing Disc. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Yu-ichiro Nakajima A Co-culture Model to Study the Effect of Kidney Fibroblast-p90RSK on Epithelial Cell Survival . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Ling Lin, Samantha White, and Kebin Hu Calcium Fluorescence Recordings from Neuroepithelial Stem Cells. . . . . . . . . . . . . . . . Masayuki Yamashita Ultrastructural Analysis of Cell–Cell Interactions in Drosophila Ovary . . . . . . . . . . . . . Matthew Antel, Valentina Baena, Mark Terasaki, and Mayu Inaba TIRF Microscopy as a Tool to Determine Exosome Composition . . . . . . . . . . . . . . . . . Noa B. Martı´n-Co freces, Daniel Torralba, Marta Lozano-Prieto, Nieves Ferna´ndez-Gallego, and Francisco Sa´nchez-Madrid Rapid Visualization of Intracellular Vesicle Events During Synaptic Stimulation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Noa B. Martı´n-Co freces, Amelia Rojas-Gomez, Sara G. Dosil, Irene Fernandez-Delgado, and Francisco Sa´nchez-Madrid Monitoring of Active Notch Signaling in Mouse Bladder Urothelium . . . . . . . . . . . . . Panagiotis Karakaidos and Theodoros Rampias Examining Local Cell-to-Cell Signalling in the Kidney Using ATP Biosensing . . . . . . Gareth W. Price, Joe A. Potter, Bethany M. Williams, Chelsy L. Cliff, Mark J. Wall, Claire E. Hills, and Paul E. Squires Isolation and Assessment of Pancreatic Islets Versus Dispersed Beta Cells: A Straightforward Approach to Examine Cell–Cell Communication . . . . . . . . . . . . . . . Rachel T. Scarl, William J. Koch, Kathryn L. Corbin, and Craig S. Nunemaker

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Contents

Promoter Pull-Down Assay: A Biochemical Screen for DNA-Binding Proteins . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Ryan R. Chaparian and Julia C. van Kessel Purification of the Vibrio Quorum-Sensing Transcription Factors LuxR, HapR, and SmcR. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Jane D. Newman and Julia C. van Kessel Preserving Cytonemes for Immunocytochemistry of Cultured Adherent Cells . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Sally Rogers and Steffen Scholpp Fluorescent Labeling of Connexin with As Complex and X-Y Coordinate Registration of Target Single Cells Based on a Triangle Standard Chip for the Image Analysis of Gap Junctional Communication. . . . . . . . . . . . . . . . . . . . . . . . Mikako Saito Chemical and Voltage Gating of Gap Junction Channels Expressed in Xenopus Oocytes . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Camillo Peracchia The Analysis of Gap Junctional Intercellular Communication Among Osteocytes in Chick Calvariae by Fluorescence Recovery After Photobleaching . . . . . Ziyi Wang, Yoshihito Ishihara, and Hiroshi Kamioka Flow Cytometry Evaluation of Gap Junction-Mediated Intercellular Communication Between Cytotoxic T Cells and Target Tumor Cells . . . . . . . . . . . . . . Mariela Navarrete, Flavio Salazar-Onfray, and Andre´s Tittarelli Index . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

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Contributors MATTHEW ANTEL • Department of Cell Biology, University of Connecticut Health Center, Farmington, CT, USA VALENTINA BAENA • Department of Cell Biology, University of Connecticut Health Center, Farmington, CT, USA RYAN R. CHAPARIAN • Biology Department, Indiana University, Bloomington, IN, USA CHELSY L. CLIFF • Joseph Banks Laboratories, School of Life Sciences, University of Lincoln, Lincoln, UK KATHRYN L. CORBIN • Department of Biomedical Sciences Heritage College of Osteopathic Medicine, Ohio University, Athens, OH, USA; Diabetes Institute, Heritage College of Osteopathic Medicine, Ohio University, Athens, OH, USA XIAOJING CUI • Department of Toxicology and Cancer Biology, University of Kentucky, Lexington, KY, USA MIGUEL F. DIAZ • Department of Integrative Biology and Pharmacology, McGovern Medical School, University of Texas Health Science Center at Houston, Houston, TX, USA; Center for Stem Cell and Regenerative Medicine, The Brown Foundation Institute of Molecular Medicine, University of Texas Health Science Center at Houston, Houston, TX, USA SARA G. DOSIL • Servicio de Inmunologı´a, Hospital Universitario de la Princesa, Universidad Autonoma de Madrid, Instituto Investigacion Sanitaria Princesa (IIS-IP), Madrid, Spain; Vascular Pathophysiology Area, Centro Nacional Investigaciones Cardiovasculares (CNIC), Madrid, Spain MIRJANA EFREMOVA • Barts Cancer Institute, Queen Mary University of London, London, UK; Wellcome Sanger Institute, Cambridgeshire, UK IRENE FERNANDEZ-DELGADO • Servicio de Inmunologı´a, Hospital Universitario de la Princesa, Universidad Autonoma de Madrid, Instituto Investigacion Sanitaria Princesa (IIS-IP), Madrid, Spain; Vascular Pathophysiology Area, Centro Nacional Investigaciones Cardiovasculares (CNIC), Madrid, Spain NIEVES FERNA´NDEZ-GALLEGO • Servicio de Inmunologı´a, Hospital Universitario de la Princesa, Universidad Autonoma de Madrid, Instituto Investigacion Sanitaria Princesa (IIS-IP), Madrid, Spain; Vascular Pathophysiology Area, Centro Nacional Investigaciones Cardiovasculares (CNIC), Madrid, Spain CLAIRE E. HILLS • Joseph Banks Laboratories, School of Life Sciences, University of Lincoln, Lincoln, UK KEBIN HU • Department of Medicine, The Pennsylvania State University College of Medicine, Hershey, PA, USA; Department of Cellular and Molecular Physiology, The Pennsylvania State University College of Medicine, Hershey, PA, USA MAYU INABA • Department of Cell Biology, University of Connecticut Health Center, Farmington, CT, USA YOSHIHITO ISHIHARA • Department of Orthodontics, Okayama University Graduate School of Medicine, Dentistry, and Pharmaceutical Sciences, Okayama, Japan HIROSHI KAMIOKA • Department of Orthodontics, Okayama University Graduate School of Medicine, Dentistry, and Pharmaceutical Sciences, Okayama, Japan PANAGIOTIS KARAKAIDOS • Biomedical Research Foundation of the Academy of Athens, Athens, Greece

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Contributors

WILLIAM J. KOCH • Department of Biomedical Sciences Heritage College of Osteopathic Medicine, Ohio University, Athens, OH, USA; Translational Biomedical Sciences, Heritage College of Osteopathic Medicine, Ohio University, Athens, OH, USA; Diabetes Institute, Heritage College of Osteopathic Medicine, Ohio University, Athens, OH, USA YOONJOO K. LEE • Departments of Pharmacology and Biochemistry, Boston University School of Medicine, Boston, MA, USA YING LIANG • Department of Toxicology and Cancer Biology, University of Kentucky, Lexington, KY, USA LING LIN • Department of Medicine, The Pennsylvania State University College of Medicine, Hershey, PA, USA; Department of Cellular and Molecular Physiology, The Pennsylvania State University College of Medicine, Hershey, PA, USA MARTA LOZANO-PRIETO • Servicio de Inmunologı´a, Hospital Universitario de la Princesa, Universidad Autonoma de Madrid, Instituto Investigacion Sanitaria Princesa (IIS-IP), Madrid, Spain; Vascular Pathophysiology Area, Centro Nacional Investigaciones Cardiovasculares (CNIC), Madrid, Spain NOA B. MARTI´N-CO´FRECES • Servicio de Inmunologı´a, Hospital Universitario de la Princesa, Universidad Autonoma de Madrid, Instituto Investigacion Sanitaria Princesa (IIS-IP), Madrid, Spain; Vascular Pathophysiology Area, Centro Nacional Investigaciones Cardiovasculares (CNIC), Madrid, Spain; Centro de Investigacion Biome´dica en Red Cardiovascular (CIBERCV), Madrid, Spain AMINA MOHAMMADALIPOUR • Department of Integrative Biology and Pharmacology, McGovern Medical School, University of Texas Health Science Center at Houston, Houston, TX, USA YU-ICHIRO NAKAJIMA • Frontier Research Institute for Interdisciplinary Sciences, Tohoku University, Sendai, Japan; Graduate School of Life Sciences, Tohoku University, Sendai, Japan MARIELA NAVARRETE • Millennium Institute on Immunology and Immunotherapy, Faculty of Medicine, Universidad de Chile, Santiago, Chile; Disciplinary Program of Immunology, Institute of Biomedical Sciences, Faculty of Medicine, Universidad de Chile, Santiago, Chile JANE D. NEWMAN • Biology Department, Indiana University, Bloomington, IN, USA CRAIG S. NUNEMAKER • Department of Biomedical Sciences Heritage College of Osteopathic Medicine, Ohio University, Athens, OH, USA; Diabetes Institute, Heritage College of Osteopathic Medicine, Ohio University, Athens, OH, USA SUMEDHA PAREEK • Immunology Program, MD Anderson Cancer Center UTHealth Graduate School of Biomedical Sciences, Houston, TX, USA CAMILLO PERACCHIA • Department of Pharmacology and Physiology, School of Medicine and Dentistry, University of Rochester, Rochester, NY, USA JOE A. POTTER • Joseph Banks Laboratories, School of Life Sciences, University of Lincoln, Lincoln, UK GARETH W. PRICE • Joseph Banks Laboratories, School of Life Sciences, University of Lincoln, Lincoln, UK THEODOROS RAMPIAS • Biomedical Research Foundation of the Academy of Athens, Athens, Greece SALLY ROGERS • Living Systems Institute, School of Biosciences, College of Life and Environmental Sciences, University of Exeter, Exeter, UK AMELIA ROJAS-GOMEZ • Servicio de Inmunologı´a, Hospital Universitario de la Princesa, Universidad Autonoma de Madrid, Instituto Investigacion Sanitaria Princesa (IIS-IP),

Contributors

xi

Madrid, Spain; Vascular Pathophysiology Area, Centro Nacional Investigaciones Cardiovasculares (CNIC), Madrid, Spain MIKAKO SAITO • Department of Biotechnology and Life Science, Tokyo University of Agriculture and Technology, Tokyo, Japan FLAVIO SALAZAR-ONFRAY • Millennium Institute on Immunology and Immunotherapy, Faculty of Medicine, Universidad de Chile, Santiago, Chile; Disciplinary Program of Immunology, Institute of Biomedical Sciences, Faculty of Medicine, Universidad de Chile, Santiago, Chile FRANCISCO SA´NCHEZ-MADRID • Servicio de Inmunologı´a, Hospital Universitario de la Princesa, Universidad Autonoma de Madrid, Instituto Investigacion Sanitaria Princesa (IIS-IP), Madrid, Spain; Vascular Pathophysiology Area, Centro Nacional Investigaciones Cardiovasculares (CNIC), Madrid, Spain; Centro de Investigacion Biome´dica en Red Cardiovascular (CIBERCV), Madrid, Spain RACHEL T. SCARL • Department of Biomedical Sciences Heritage College of Osteopathic Medicine, Ohio University, Athens, OH, USA STEFFEN SCHOLPP • Living Systems Institute, School of Biosciences, College of Life and Environmental Sciences, University of Exeter, Exeter, UK KRISTEN L. SEGARS • Departments of Pharmacology and Biochemistry, Boston University School of Medicine, Boston, MA, USA PAUL E. SQUIRES • Joseph Banks Laboratories, School of Life Sciences, University of Lincoln, Lincoln, UK MARK TERASAKI • Department of Cell Biology, University of Connecticut Health Center, Farmington, CT, USA ANDRE´S TITTARELLI • Programa Institucional de Fomento a la Investigacion, Desarrollo e Innovacion (PIDi), Universidad Tecnologica Metropolitana (UTEM), Santiago, Chile DANIEL TORRALBA • Servicio de Inmunologı´a, Hospital Universitario de la Princesa, Universidad Autonoma de Madrid, Instituto Investigacion Sanitaria Princesa (IIS-IP), Madrid, Spain; Vascular Pathophysiology Area, Centro Nacional Investigaciones Cardiovasculares (CNIC), Madrid, Spain VICKERY TRINKAUS-RANDALL • Departments of Pharmacology and Biochemistry, Boston University School of Medicine, Boston, MA, USA JULIA C. VAN KESSEL • Biology Department, Indiana University, Bloomington, IN, USA ROSER VENTO-TORMO • Wellcome Sanger Institute, Cambridgeshire, UK MARK J. WALL • School of Biomedical Science, University of Warwick, Coventry, UK ZIYI WANG • Department of Orthodontics, Okayama University Graduate School of Medicine, Dentistry, and Pharmaceutical Sciences, Okayama, Japan; Research Fellow of Japan Society for the Promotion of Science, Tokyo, Japan PAMELA L. WENZEL • Department of Integrative Biology and Pharmacology, McGovern Medical School, University of Texas Health Science Center at Houston, Houston, TX, USA; Center for Stem Cell and Regenerative Medicine, The Brown Foundation Institute of Molecular Medicine, University of Texas Health Science Center at Houston, Houston, TX, USA; Immunology Program, MD Anderson Cancer Center UTHealth Graduate School of Biomedical Sciences, Houston, TX, USA SAMANTHA WHITE • Department of Medicine, The Pennsylvania State University College of Medicine, Hershey, PA, USA; Department of Cellular and Molecular Physiology, The Pennsylvania State University College of Medicine, Hershey, PA, USA BETHANY M. WILLIAMS • Joseph Banks Laboratories, School of Life Sciences, University of Lincoln, Lincoln, UK

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Contributors

MASAYUKI YAMASHITA • Center for Basic Medical Research, International University of Health and Welfare, Ohtawara, Japan CUIPING ZHANG • Department of Toxicology and Cancer Biology, University of Kentucky, Lexington, KY, USA XINGHUI ZHAO • Department of Toxicology and Cancer Biology, University of Kentucky, Lexington, KY, USA

Methods in Molecular Biology (2021) 2346: 1–10 DOI 10.1007/7651_2020_343 © Springer Science+Business Media, LLC 2021 Published online: 25 February 2021

Inference of Ligand–Receptor Pairs from Single-Cell Transcriptomics Data Mirjana Efremova and Roser Vento-Tormo Abstract Cell–cell communication is crucial for development and tissue homeostasis in multicellular organisms. Single-cell transcriptomics has emerged as a revolutionary technique for dissecting cellular compositions and potential cell–cell communication events via ligand–receptor pairs. To provide a systematic characterization of intercellular communication, we developed a framework to map cell–cell communication events mediated by ligand–receptor interactions across different cell types using single-cell transcriptomics data. Our repository of ligands, receptors and their interactions is integrated with a computational approach to identify cell-type specific and biologically relevant interactions. Here, we summarize the structure and content of our repository and present a practical guide for inferring cell–cell communication networks from single-cell RNA sequencing data. Key words Cell–cell communication, Single-cell RNA sequencing, Receptors, Ligands

1

Introduction Intercellular communication underlies numerous biological processes including coordination of cellular fate, behavior, and response to neighboring cells, in healthy and diseased conditions. Single-cell transcriptomics, by measuring the expression of ligands and receptors in individual cell types, has provided powerful means for mapping interactions and inferring cellular communication networks [1, 2]. To provide a systematic characterization of cell–cell communication, we developed CellPhoneDB, a public repository of ligands, receptors and their interactions, integrated with a statistical framework to identify interactions enriched on specific cell types [2, 3]. Our database relies on public resources as well as manual curation and takes into account the subunit architecture of ligands and receptors, to accurately represent heteromeric complexes (see Notes 1 and 2). To infer intercellular communication networks between different cell types, we consider the expression levels of ligands and receptors within each cell type, and use empirical shuffling to

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calculate which ligand–receptor pairs display significant cell-type specificity. Briefly, we randomly permute the cluster labels of all cells and calculate the mean of the average ligand expression level in a cell type and the average receptor expression level in the interacting cell type, generating a null distribution. By calculating the proportion of the means which are as or higher than the actual mean, we obtain a p-value for the likelihood of cell-type specificity of each ligand–receptor complex. In this way, our method predicts biologically relevant molecular interactions that can be visualized using intuitive tables and plots. Our tool is available as a user-friendly interface at cellphonedb. org and as a python package on github (https://github.com/ Teichlab/cellphonedb).

2

2.1

Materials

Input Data Files

CellPhoneDB can be used either through the interactive website (https://www.cellphonedb.org/) or as a Python package using the user’s computer/cloud/farm. The Python package is recommended for large datasets (datasets larger than 10 GB). – Metadata input file: cell type annotation file containing cluster annotation of each cell. Clustering can be performed with standard packages for scRNA-seq analysis such as Seurat [4], SCANPY [5]. The metadata file has two columns: “Cell”, which represents the name of the cell, and “cell_type”, which represents the name of the cluster. Accepted formats are .csv (for comma separated); .txt, .tsv, .tab (for tab separated); and pickle. – Counts input file: single-cell RNA sequencing (scRNA-seq) count data with gene expression values where cells are in columns are cells and genes are in rows. Accepted gene names identifiers include Ensembl IDs, gene names and hgnc_symbol annotations. Use of normalized count data is recommended. Accepted formats are .csv (for comma separated); .txt, .tsv, .tab (for tab separated); and pickle (see Notes 3 and 4).

2.2

Software

– Python 3.5 or higher. – SQLAlchemy (see Note 5). – SQLite.

2.3

Hardware

– Linux or MAC OS.

Inference of Ligand–Receptor Pairs from Single-Cell Transcriptomics Data

3 3.1

3

Methods Installation

We recommend using a virtual environment, but this is optional (step 1). 1. Create and activate a python virtual environment. python -m venv cpdb-venv source cpdb-venv/bin/activate

2. Install CellPhoneDB. pip install cellphonedb

3.2 Running in the Statistical Analysis Mode

Run CellPhoneDB with statistical analysis using the input file names (specifying the full path to the files). cellphonedb method statistical_analysis usermetafile.txt usercountsfile.txt

Optional parameters: --project-name: Project’s name. A subfolder with this identity is created in the output folder. ./out is used by default. --iterations: Number of iterations for statistical analysis. 1000 interactions are used by default. --threshold: % of cells expressing the specific ligand or receptor --result-precision: Number of decimal digits in results. 3 decimal digits are used by default. --counts-data: [ensembl | gene_name | hgnc_symbol] Gene identifiers used in the counts data --output-path: Directory where the results will be allocated (the directory must exist) ./out is used by default. --output-format: Output format of the results files (extension will be added to filename if not present). txt is used by default. --means-result-name: Name of the means result file. means.txt is used by default. --significant-mean-result-name: Name of the significant means result file. Significant_means.txt is used by default --deconvoluted-result-name: Name of the deconvoluted result file. deconvoluted.txt is used by default --verbose/--quiet: Print or hide cellphonedb logs [verbose] --pvalues-result-name: Name of the pvalues result file. pvalues.txt is used by default --debug-seed: Debug random seed -1. To disable it please use a value >=0 . -1 is used by default --threads: Number of threads to use. >=1. 4 is used by default.

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l

A usage example to set output path, number of iterations, and number of threads: mkdir custom_folder cellphonedb method statistical_analysis usermetafile.txt usercountsfile.txt --outputpath=custom_folder --iterations=10 --threads=2

3.3 Subsampling and Statistical Analysis

To improve the speed and efficiency and reduce memory requirements when analyzing large datasets, we implemented a subsampling algorithm [6] in our framework. When using the subsampling option, the user needs to specify whether the input count data was log-transformed. Run CellPhoneDB with statistical analysis using the input files and add --subsampling with subsampling-specific parameters. cellphonedb method statistical_analysis usermetafile.txt usercountsfile.txt -subsampling --subsampling-log true

In addition to the parameters described in step 3, there are also subsampling specific parameters. --subsampling-log: Enable log transformation for non-log transformed data inputs. This is a mandatory parameter. --subsampling-num-pc: Subsampling NumPC argument --subsampling-num-cells: Number of cells to subsample to. 1/3 of the cells is used by default.

3.4 Normal Mode, Without Statistical Analysis

Run CellPhoneDB without statistical analysis using the input files and a defined --threshold parameter. The parameters are the same as described in step 3, with the exception of --pvalues-result-name, --threads, and --debug-seed which are specific for the statistical analysis and should not be used here. cellphonedb method analysis usermetafile.txt usercountsfile.txt

3.5

Visualization

The user can visualize the analysis results using dot plots and heatmaps. 1. For dot plot visualization, run the dot plot visualization command using the means.csv and pvalues.scv output files. This visualization is only relevant for the statistical analysis mode. cellphonedb plot dot_plot

Inference of Ligand–Receptor Pairs from Single-Cell Transcriptomics Data

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Dot plot specific parameters: --means-path: The means output file. ./out/means.txt is used by default. --pvalues-path: The pvalues output file ./out/pvalues.txt is used by default. --output-path: Output folder. ./out is used by default. --output-name: Name of the output plot. plot.pdf is used by default. Available output formats are restricted to those supported by R’s ggplot2 package (e.g. pdf, png, jpeg) --rows: File with a list of rows to plot, one per line --columns: File with a list of columns to plot, one per line --verbose / --quiet: Print or hide cellphonedb logs [verbose]

To plot only specific rows/columns, use text files with a list of interactions/celltype–celltype comparisons (each interaction/comparison in a new row). cellphonedb plot dot_plot --rows rows.txt --columns columns.txt

2. For heatmap visualization, run the heatmap visualization command, using the pvalues.scv output file. This visualization is only relevant for the statistical analysis mode. cellphonedb plot heatmap_plot meta_data

Specific parameters for the heatmap plot: --pvalues-path: The pvalues output file. ./out/pvalues.txt is used by default. --output-path: Output folder. default: ./out is used by default. --count-name: Filename of the output plot heatmap_count.pdf is used by default. --log-name: Filename of the output plot using logcount of interactions. heatmap_log_count.pdf is used by default --count-network-name: Filename of the output network file. network.txt is used by default. --interaction-count-name:

Filename

of

the

output

interactions-count file. interaction_count.txt is used by default. --verbose / --quiet: Print or hide cellphonedb logs [verbose]

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3.6 Different Versions of the CellPhoneDB Database

For reproducibility of their analysis, the users have access to different versions of the CellPhoneDB databases and they can specify which version they prefer to use. The users can list and download available versions of the database from a remote repository (the CellPhoneDB official available data). 1. Use a specific database version by adding the parameter -database to the command “cellphonedb method”. cellphonedb

method

statistical_analysis

usermetafile.txt usercountsfile.txt --database=v0.0.4

The –database parameter can be either a database file, in which case it will be used as it is, or a version number, in which case the version that matches the specified version parameter will be used. If the specified database version does not exist locally on the user’s computer it will be downloaded from the remote repository. If the --database argument is not specified, the latest available local database version will be used for the analysis. The downloaded database versions will be stored in a folder under ~/.cpdb/releases. 2. List available database versions from the remote CellPhoneDB repository. cellphonedb

database

list_remote

3. List available versions from the local repository, stored locally on the user’s computer. cellphonedb

database

list_local

4. Download a specific version from the remote repository. cellphonedb

database

download

or cellphonedb database download --version

version_spec must be one of the database versions in the database listed by the list_remote command. The latest available version will be downloaded by default. This can be done also by using latest as a version_spec. 3.7 Generating a User-Specific Database

Users can also create a custom database, using their own input files. They can choose whether to merge their database with the current CellPhoneDB input files or to only use their own database (see Note 6). Generate a custom database with user-specific input files. cellphonedb database generate

Inference of Ligand–Receptor Pairs from Single-Cell Transcriptomics Data

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Database generate specific parameters: --user-protein: Protein input file --user-gene: Gene input file --user-complex: Complex input file --user-interactions: Interactions input file --fetch: Specific lists can be downloaded from original sources while creating the database, e.g., uniprot, ensembl. The input tables included in the CellPhoneDB package will be used by default. To enable downloading an updated copy from the remote servers --fetch must be appended to the command --result-path: Output folder --log-file: Log file

The database file will be generated in the --result-path folder (by default in the folder “out” if an output folder is not specified) with cellphonedb_user_{datetime}.db. The user defined input tables will be by default merged with the current CellPhoneDB input tables. To use this database, use the --database parameter when running the “cellphonedb method” command: cellphonedb method statistical_analysis in/example_data/test_meta.txt in/example_data/test_counts.txt --database out/cellphonedb_user_2019-05-10-11_10.db l

Use only user-specific interactions without merging with the current CellPhoneDB database (using custom_interaction_file. csv as a comma separated input file with a list of interactions— with mandatory columns as described in [3]). cellphonedb database generate --userinteractions custom_interaction_file.csv --user-interactions-only

l

Add or modify some existing interactions (using custom_interaction_file.csv as a comma separated input file with a list of interactions to add/modify—with mandatory columns as described in [3]): cellphonedb database generate --user-interactions custom_interaction_file.csv

l

For duplicated interactions, the user specified interactions will overwrite the CellPhoneDB interactions. Modify any protein data (using custom_protein_file.csv as a comma separated input file with a list of proteins to overwrite—with mandatory columns as described in [3]).

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Mirjana Efremova and Roser Vento-Tormo cellphonedb database generate --user-protein custom_protein_file.csv

l

Update database from remote servers, for example from UniProt, IMEx, and ensembl. cellphonedb database generate –fetch

3.8

Help Option

Get a detailed description of the mandatory and optional parameters using the help option. cellphonedb method statistical_analysis metafile.txt countsfile.txt --help

3.9 Interactive Web Portal

The web interface includes input forms for the user to specify analysis parameters before submission. Downstream calculations are performed on the application’s servers. 1. Upload your input files in the “Exploring your scRNAseq” tab. You can get an update when the analysis is finished by providing your email address. 2. When the analysis has finished, the “significant_means” results table will appear. Results can be downloaded and the current view can be changed by clicking on the ”Data Shown” button. To display detailed information for the specific interaction pair, you can click on any field from the id_cp_interaction column. 3. To visualise the results, go to the “Plots” tab and choose the type of plot you would like to be generated. For plotting dot plots, select specific columns and rows of the interactions you are interested in. The online results viewer allows the user to select specific columns to be displayed in each table.

4

Notes 1. CellPhoneDB stores ligand–receptor interactions and information related to interacting molecules. For example, we annotated the subunit architecture of each of the interacting partners as well as multiple gene and protein identifiers. All this information is stored in four main .csv data files that are uploaded into the database: “gene_input.csv”, “protein_input. csv”, “ complex_input.csv”, and “interaction_input.csv”. The “gene_input” file stores information related to the gene nomenclature. This information is key to establishing a connection within the scRNA-seq count data and the

Inference of Ligand–Receptor Pairs from Single-Cell Transcriptomics Data

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interaction pairs stored in the database as proteins. Specifically, it stores: (a) gene name (“gene_name”), (b) UniProt identifier (“uniprot”), (c) HUGO nomenclature committee symbol (HGNC) (“hgnc_symbol”), and (d) gene ensembl identifier (ENSG) (“ensembl”). The “protein_input” file stores the identity of proteins and combines manual annotation of the protein identities (based on bibliography or uniprot description) as well as data downloaded directly from uniprot. The “complex_input” file stores the identity of each of the multisubunit proteins that will form the heteromeric ligands and receptors. A specific name is given to each heteromeric complex. Finally, the “interaction_input” file contains the protein–protein interactions stored in CellPhoneDB. For binary interactions, each interaction partner is annotated by their UniProt identifier. For interactions involving heteromers, the name of the complex is introduced. The identity of the complex is defined in “complex_input”. Each interaction has a unique identifier (“id_cp_interaction”) generated automatically by the pipeline. For further details on the content of the tables, please refer to [3]. 2. CellPhoneDB is a curated database, which means that only information supported by bibliography is stored. This information is reviewed internally by members of our team and therefore, could be biased toward specific expertise (e.g., immunology, development). We encourage users sending interacting proteins involved in cell–cell communication, together with a DOI of the publication supporting this information. Information can be shared by e-mail, the form available at www.cellphonedb.org or pull request to the CellPhoneDB data repository (https://github.com/Teichlab/cellphonedb-data). All the information will be checked by one of our team members and uploaded into the database. 3. CellPhoneDB was created for scRNA-seq but can be used for other types of data, for example bulk RNA-seq data from pure cell populations. 4. CellphoneDB was created using human-specific ligand–receptor interactions, but it can also be applied on datasets from other species by using orthologs. 5. CellPhoneDB data is stored in an SQLite relational database (https://www.sqlite.org/). The structure and logic of the database was built using SQLAlchemy (www.sqlalchemy.org) and Python 3. All the code is open source and available through https://github.com/Teichlab/cellphonedb. For further guidance into the internal structure of the database, please refer to [3].

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6. CellPhoneDB users can also generate their own cell–cell interactions files using our github tool (https://github.com/Teichlab/cellphonedb). For the lists to work in our database, the format of the users’ lists must be compatible with the input files of our database. Please refer to [3] for further information.

Acknowledgments We thank Sarah Teichmann for useful guidance on the method and curation of protein–protein interactions; Gavin J. Wright, Laura Wood, and Gerard Graham for advice on protein–protein interactions; and Miquel Vento-Tormo and YDEVS members for their help with the webserver and the implementation of the code in github. M.E. is funded by a Barts Charity Lectureship (grant MGU045). The project was supported by Wellcome Sanger core funding (no. WT206194) and a Wellcome Strategic Support Science award (no. 211276/Z/18/Z). References 1. Camp JG et al (2017) Multilineage communication regulates human liver bud development from pluripotency. Nature 546:533–538 2. Vento-Tormo R et al (2018) Single-cell reconstruction of the early maternal-fetal interface in humans. Nature 563:347–353 3. Efremova M, Vento-Tormo M, Teichmann SA, Vento-Tormo R (2020) CellPhoneDB: inferring cell-cell communication from combined expression of multi-subunit ligand-receptor complexes. Nat Protoc 15:1484–1506

4. Satija R, Farrell JA, Gennert D, Schier AF, Regev A (2015) Spatial reconstruction of single-cell gene expression data. Nat Biotechnol 33:495–502 5. Wolf FA, Angerer P, Theis FJ (2018) SCANPY: large-scale single-cell gene expression data analysis. Genome Biol 19:15 6. Hie B, Cho H, DeMeo B, Bryson B, Berger B (2019) Geometric sketching compactly summarizes the single-cell transcriptomic landscape. Cell Syst 8:483–493

Methods in Molecular Biology (2021) 2346: 11–20 DOI 10.1007/7651_2020_329 © Springer Science+Business Media New York 2020 Published online: 07 November 2020

Multiple Imaging Modalities for Cell-Cell Communication via Calcium Mobilizations in Corneal Epithelial Cells Yoonjoo K. Lee, Kristen L. Segars, and Vickery Trinkaus-Randall Abstract Chemical indicators are used to study calcium signaling events in the context of live cell imaging. Fluo-3 AM, Fluo-4 AM, and Cal-520 AM are three commonly used fluorescent indicators derived from the calcium chelator BAPTA. Here we describe sample protocols that detail how these indicators are used in in vitro and ex vivo experiments to analyze the role of calcium mobilizations in cell-cell communication and coordinated cellular motility in the context of wound healing. Key words Ex vivo in situ analysis, Confocal imaging, Calcium mobilization, Cell-cell communication, Live cell imaging, Image analysis

1

Introduction What do neurotransmitter release, skeletal muscle contraction, and signal transduction of Gq-type G protein-coupled receptors (GPCRs) have in common? These processes all take advantage of changes in the intracellular concentration of calcium ions to cause changes in the cell. In the cytosol, the concentration of calcium ions is tightly maintained at 100 nM, roughly 20,000–100,000-fold lower than the concentration in the extracellular space [1]. Due to the role of calcium ions in regulating enzyme and protein activity, small alterations in this concentration can cause large changes in gene transcription, protein activity, and cellular function. In muscles, calcium released from the sarcoplasmic reticulum binds to troponin [1–3]. This causes a change in its 3D configuration that allows binding sites on actin proteins to become exposed [1– 3]. Myosin is now able to bind to this site and hydrolyze ATP to cause the sarcomere to contract [1–3]. In neurons, activation of voltage-gated Ca channels causes a sudden increase in the calcium concentration at the axon terminal [4]. This change prompts the

Yoonjoo Lee and Kristen Segars are co-first authors. Electronic Supplementary Material: The online version of this chapter (https://doi.org/10.1007/7651_ 2020_329) contains supplementary material, which is available to authorized users.

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release of neurotransmitter-filled vesicles into the synaptic cleft, propagating the action potential to an adjacent cell [4]. Signaling through Gq-type GPCRs occurs by cleaving the membrane lipid phosphatidylinositol biphosphate (PIP2) to inositol triphosphate (IP3) and diacylglycerol (DAG) upon ligand binding [1]. IP3 binds to a receptor on the endoplasmic reticulum, causing the release of stored calcium into the cytosol [1]. This calcium binds to and activates many different proteins resulting in changes in transcription and translation. These are just three examples, but there are countless other instances where calcium signaling affects cellular function and cell-cell communication. Given the involvement of calcium signaling in such a diverse range of processes, having a reliable way to monitor and analyze calcium mobilization events is of great interest to researchers in many different fields. Calcium indicators respond to changes in cytosolic calcium concentration with changes in their fluorescent properties [5]. They can be grouped into two main classes: genetically encoded and chemical [5]. Genetically encoded indicators are often derived from GFP or its variants fused with calmodulin and the M13 domain of myosin light chain kinase [5]. These indicators have the advantage of localization to particular cellular subpopulations and can yield data from in vivo systems such as that following calcium mobilizations in neurons [5]. These indicators have been used to map the role of anesthetics on the response of specific motor neurons in C. elegans [6]. Although extremely useful in many contexts, we will not be discussing them further in this chapter. Like genetically encoded calcium indicators, chemical indicators exhibit substantial changes in fluorescence with increases in cytosolic calcium concentration [5]. However, there are several key differences between the two. Whereas genetically encoded indicators are incorporated into an organism’s genome and allow the organism to produce the fluorescent protein itself, chemical indicators are exogenous substances that are preincubated with ex vivo or in vitro samples [5] (see Note 1). The Fura and Indo dyes require UV lasers for excitation and are ratiometric dyes that demonstrate specific changes in maximum excitation or emission wavelengths, respectively, upon changes in intracellular calcium concentration [5, 7]. The Fluo series do not require UV lasers and exhibit greater fluorescence between bound and unbound forms, but they do lack the shift in emission spectra observed in the other probes so that one must carefully decide which probe to use [7]. The Fluo chemical indicators are based on the nonfluorescent calcium chelator 1,2-bis(o-aminophenoxy) ethane-N,N,N0 ,N00 -tetraacetic acid (BAPTA) [7]. BAPTA itself is structurally similar to the metal ion chelator ethylenediaminetetraacetic acid (EDTA) [7]. BAPTA itself is structurally similar to the metal ion chelator EDTA but has been modified to yield a higher

Live Cell Imaging

13

affinity for Ca2+ ions and a lower affinity for other metal ions such as Fe3+ and Mg2+ [7]. The indicators have been modified to optimize affinity for calcium and yield maximum fluorescence at desired wavelengths [8]. When calcium binds to an indicator, it undergoes changes in its 3D structure that greatly enhance its fluorescence, up to 100-fold in the case of Fluo-3 [7]. In this method discussion, we are focusing on three of these chemical indicators: Fluo-3 AM, Fluo-4 AM, and Cal-520 AM. Fluo-3 AM and Fluo-4 AM are engineered to fluoresce upon the structural changes induced upon Ca ion binding [5, 7, 8]. Fluo4 is a variant of Fluo-3 that has replaced two chlorine atoms with fluorine [5, 7, 8]. This results in a higher fluorescent excitation at a wavelength of 488 nm and a higher overall signal [8]. Their maximum excitation/emission spectra are 506/526 and 494/516, respectively [8]. Advantages of Fluo-4 include faster loading and a higher signal at the same concentration [8]. Both Fluo-3 and Fluo4 are attached to an acetoxymethyl ester group that blocks the carboxyl groups on these molecules and facilitates crossing the plasma membrane [5, 7, 8]. This moiety is cleaved by esterases in the cell, exposing the carboxyl groups and preventing diffusion out of the cell [5, 7, 8]. The indicators are denoted as Fluo-3 AM or Fluo-4 AM if they have this moiety. In addition to Fluo-3 AM and Fluo-4 AM, there are two additional indicators in the Fluo series: Fluo-5 AM and Fluo-8 AM [8]. Fluo-5 AM is structurally similar to Fluo-4 AM but exhibits lesser affinity for calcium ions [7]. This makes Fluo-5 AM suitable for detecting calcium concentrations between 1 μM and 1 mM, concentrations that would saturate the response of Fluo-3 AM or Fluo-4 AM [7]. Fluo-8 AM was engineered to facilitate loading while maintaining the same useful excitation and emission spectra [9]. Whereas Fluo-3 AM and Fluo-4 AM must be loaded at 37  C, Fluo-8 AM can be loaded at room temperature [9]. Fluo-8 AM is also twice as bright as Fluo-4 AM and four times as bright as Fluo-3 AM [9]. The Fluo series are useful for short-term calcium imaging ( 150,000. 6. Coverslips for immunofluorescence (IF) (thickness No. 1.5H; 0.170 mm  0.005 mm such as 0117530 Marienfeld). 7. Coverslip-bottom dishes for imaging. Use commercial (35 mm diameter MatTek Corporation, iBIDI) or homemade chambers no. 1.5 thickness to optimize imaging (see Note 2). 8. Isolation of exosomes: ultracentrifuge tubes 39 ml, clear; thin walls. 9. Western blot analysis: 5 protein-loading buffer, non-reducing 10–12% SDS-PAGE, nitrocellulose (0.20 or 0.45 μm pore size) or PVDF membranes. 10. Protein quantification: BCA or Bradford assay kit, 2 mg/ml BSA stock for standard curve, 96-well flat-bottom plates, spectrophotometer. 11. Petri dishes for PBMC adhesion (peripheral blood mononuclear cells). 12. Flasks for cell culture (different sizes).

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2.3 Media and Solutions/Buffers

1. Complete medium: RPMI 1640 supplemented with HEPES (25 mM), nonessential amino acids, glutamine (100 mM), FCS (fetal calf serum, heat inactivated (55  C, 30 min; 10%), FCS will be ultracentrifuged for exosome production and recovery (see Note 3). Cell media for mouse cells is also supplemented with β-mercaptoethanol (1 mM). 2. Incomplete medium: RPMI 1640, HEPES (25 mM), L-glutamine (100 mM), nonessential amino acids. 3. Wash solution: Hanks’ Balanced Salt Medium (HBSS). 4. Saline solution: NaCl (154 mM). Sterile. 5. Lymphocyte separation medium: any homemade or commercial media such as Ficoll Histopaque or Biocoll. 6. Coating buffer: bicarbonate-carbonate medium. NaHCO3 (0.1 M), Na2CO3 (0.032 M), pH: 9.6. 7. Lysis buffer for Western blot: 50 mM Tris–HCl (pH 7.4), 0.2% Triton X-100, 1% NP40, 150 mM NaCl, 1.5 mM MgCl2, 2 mM EDTA, and protease and phosphatase inhibitors. 8. TBS (Tris-buffered saline): Tris–HCl 50 mM (pH 7.4), NaCl (154 mM). 9. PHEM (2): 120 mM pipes, 50 mM HEPES, 20 mM EGTA, 4 mM MgCl2; pH 6.9. 10. Fixation solution: HBSS, 1% paraformaldehyde (PFA), 5% sucrose. We recommend the use of specific solutions for microscopy, such as RT15710 from Electron Microscopy Sciences. 11. Anti-fading solution: glycine 200 mM in Tris–HCl (pH 7.4). 12. Immunofluorescence (IF) blocking solution: 3% BSA, 20 μg/ ml human gamma-globulins in HBSS (see Note 4). 13. IF permeabilizing solution: 0.2% TX-100 solution (HBSS supplemented with 0.5% paraformaldehyde and 1.5% BSA). 14. IF blocking and permeabilizing solution: ultracentrifuged PHEM (1), BSA 3%, human γ-globulin 100 μg per ml, sodium azide 0.2%, 0.2% Triton Tx-100. 15. Water-based mounting medium: we recommend the use of a controlled medium, either homemade or commercial such as ProLong Gold Antifade Mountant (P36930 Thermo Fisher Scientific), or Citifluor Phosphate-Buffered Saline AF3 (refractive index: 1.338 at 20  C; pH 10; optically transparent from 300 to 750 nm, glycerol free; AF3 Citifluor).

2.4

Equipment

1. Ultracentrifuge: any equipment able to reach 120,000  g, with rotors for swing buckets; tubes 39 ml. 2. Nanosight NS300, Malvern Panalytical: infrared device for detection of nanoparticles in solution, 10–2000 nm range.

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3. TIRFM imaging: Leica AM TIRF MC M system mounted on a Leica DMI 6000B fitted with a HCX PL APO 100  1.46 NA oil objective microscope, coupled to an Andor-DU8285 VP-4094 camera (see Note 5). 2.5

Software

1. Nanosight NTA 2.3 software for nanoparticles 2. LAS-AF 2.6.0. Build 7,266 for image acquisition. 3. Image J 1.51n software (Wayne Rasband, National Institutes of Health, USA. http://imagej.nih.gov/ij; Java 1.8.0_66; 64 bit) image analysis 4. Imaris 7.2.2 (Bitplane) for image analysis.

3

Methods

3.1 CD4+ T-Cell Purification from Human Peripheral Blood Lymphocytes (huPBLs)

1. Isolate the PBMCs from complete blood (50–200 ml) or buffy coat preparations (60 ml from 450 ml complete blood) from healthy human donors through a Ficoll Histopaque gradient. Buffy coat should be diluted 1:2 (v/v) with saline solution before mounting gradients. Centrifuge for 700  g, 20 min, R/T without brake or acceleration. 2. Wash the cells recovered from the interphase with the Ficoll with saline solution four to six times to drain the platelets (PBMCs). 3. Purify CD3+CD4+ cells by negative selection with magnetic beads-based methods. A cocktail of antibodies and streptavidin-conjugated beads for autoMACS (Miltenyi Biotec) or Stem Cell Solutions is recommended.

3.2 Generation of Lymphoblasts from Human PBLs: PHA or SEE-Specific

1. The PBMCs are allowed to adhere in complete or incomplete medium (at least two rounds of plate adhesion) for 30 min at 37  C to deplete monocytes and granulocytes by plate adhesion (see Note 6). 2. After monocyte depletion, plate cells at 2  106 per ml in complete medium. 3. Grow cells with SEE (0.01 μg/ml) for 48–72 h (see Note 7). Alternatively, cells can be treated with phytohemagglutinin A (PHA) (1 μg/ml) for 48 h. 4. Wash SEE or PHA (1000  g, 10 min), and grow cells in complete medium supplemented with IL-2 (20–50 Units/ ml). Add IL-2 every 2 days. 5. Cells can be used at day 7 to produce exosomes. 6. SEE-treated cells can be stimulated with SEE (0.1 μg/ml) and PHA (0.4 μg/ml) for 18–24 h a week later. Wash them and grow in complete medium with IL-2 as above. Cells can be

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used upon 18–24 h of cell culture. The percentage of Vβ8+ cells could then be around 40–60% (anti-Vβ8-FITC, BD Biosciences, for flow cytometry; see Note 8). 7. Cells can be restimulated with SEE and PHA every 15 days. Lymphoblasts can be frozen (107/ml). 3.3 Isolation of EVs from Cell Culture

Leukocytes and leukocyte-related cell lines constitutively release EVs that accumulate in the cell medium. The ultracentrifuged FCS is therefore essential to isolate cell-derived exosomes (see Fig. 1).

3.3.1 Production and Isolation of EVs (Fig. 1a)

1. Perform a quick Ficoll gradient by spinning cells at 500  g, 5 min, R/T, no brake, to eliminate any dead or apoptotic cell, and remove culture medium used to amplify cells. 2. Recover live cells; wash once in clean, new HBSS. 3. Culture cells at 1–3  106 cells per ml in RPMI, 10% ultracentrifuged FCS for 48–72 h at 37  C, 5% CO2 (see Note 9). 4. To isolate EVs from the conditioned medium, remove cells by centrifugation at 320  g for 5 min, and collect the supernatant. 5. Spin the supernatant at 2000  g, 20 min, 4  C, and collect the supernatant. The pellet contains cell debris. 6. Spin the supernatant at 10,000  g, 30 min, 4  C, and collect the supernatant. The pellet contains possible organelle fragments and large vesicles. Use maximal acceleration rate. Stop centrifugation using either a low braking rate or not brake at all. 7. Spin the supernatant at 100,000  g, 70 min, 4  C, and collect the EVs (i.e., exosomes) in the pellet (low brake). 8. Aspirate carefully the supernatant, and resuspend all the pellets pertaining to similar experimental conditions in HBSS to wash them together once. 9. Centrifuge at 100,000  g, 70 min, 4  C. Discard the supernatant by aspirating carefully (low brake). 10. Resuspend the pellet in less than 300 μl of ultraclean (i.e., ultracentrifuged) HBSS. Some media traces may remain after the former aspiration step; adjust volumes so that final volume is similar to all samples.

3.3.2 Quantification and Characterization of EVs

Isolated EVs can be quantified and characterized through different methods, such as quantification of its number, size, and biochemical enrichment in specific markers. 1. Measure exosome content through an NTA (nanoparticle tracking analysis). Keep 10 μl of the sample and assay 1:100

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Fig. 1 Workflow for exosome isolation and immunofluorescence. (a) Cells are cultured in ultracentrifuged medium for 48–72 h. The supernatant is subjected to a series of differential centrifugations, and the resulting pellet is washed and ultracentrifuged again at 100,000  g. The exosomes are ready to be quantified and stained. (b) Exosomes are centrifuged in a sucrose cushion and then allowed to stick gently to the surface of the poly-L-Lysine treated coverslip. Once attached, they are ready to continue with the staining protocol. (c) Exosomes are incubated in a coverslip coated with antibodies against exosome markers. After two washes, only the vesicles containing specific exosomal proteins are adhered to the coverslip and are ready to continue with the staining protocol

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Fig. 2 Graphs and data generated by Nanosight NS300. Diluted isolated exosomes (1:200 in HBSS) were imaged and processed through nanoparticle tracking analysis (NTA). Three different videos were acquired for 30 s and analyzed for average number of particles and size (S1 to S3)

and 1:200 dilutions to measure number and size of particles in solution. Sometimes higher dilutions are required (see Fig. 2 as example). 2. Quantification by BCA or Bradford assay: keep 25 μl of the sample for protein assay and use it by adding NP-40 to a final concentration of 0.5%. The calibration curve will include serial dilutions of BSA ranging from 2 mg/ml to 25 μg/ml. Prepare twofold curve and pipette duplicate aliquots of 10 μl of exosome samples into 96-well plate. Add the BCA or Bradford reagent. 3. Western blot analysis: add 12.5 μl of Laemmli solution (5) to 50 μl of the exosome sample, and boil the mix at 70  C for 10 min. Process samples by 10 or 12% non-reducing SDS-PAGE and immunoblot to probe the membranes for specific markers (e.g., tetraspanins CD63 and CD81, Tsg101), cytoskeletal proteins (α-actinin, ERMs, α-tubulin), and other proteins. Use secondary antibodies conjugated to HRS to detect the antibodies (see Note 10). 3.4 Coating of Coverslips and Coverslip-Bottom Dishes

All reagents should be ultracentrifuged or centrifuged to exclude the possibility of adding small-size particles increasing background, e.g., in 2.0 ml tubes in a benchtop centrifuge 18,000  g, 30 min, 4  C (see Note 11).

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1. Incubate the imaging surfaces with Poly-L-Lys hydrobromide in sterile water (250 μg/ml). 2. Pipette 25–50 μl of the centrifuged mix per coverslip; incubate o/n at 4  C (see Notes 2 and 12). 3. Wash twice with centrifuged sterile water. 4. Store in centrifuged HBSS until used. They can be frozen at this step. Do not allow to dry.

3.4.2 Preparation of Antibody-Coated Surfaces

1. Dilute the corresponding antibodies in coating buffer (0.1–10 μg/ml): anti-CD81, anti-CD9, anti-CD63 monoclonal antibodies, or a mix of them. 2. Pipette 25–50 μl of the mix per 10 mm coverslip or coverslipbottom dish, and incubate o/n at 4  C. 3. Wash twice with HBSS or incomplete medium. 4. Block with BSA 3% for 20 min, R/T. 5. Wash twice with HBSS or incomplete medium. 6. Store in HBSS until used. They can be frozen at this step. Do not allow to dry.

3.5 Sample Preparation for Imaging

3.5.1 Preparation of Samples on Poly-L-LysineCoated Surfaces (Fig. 1b)

All solutions used for coverslip immunofluorescence should be ultracentrifuged at 100,000  g for 70 min (4  C) in a swinging bucket rotor or centrifuged in 2.0 ml tubes in a benchtop centrifuge at 13,200  g, 30 min, 4  C in a fixed-angle rotor to exclude the possibility of adding small-sized particles to preparations increasing debris and background (see Note 11). 1. Spin around 8  106 isolated exosomal particles in a sucrose cushion (10% in HBSS; 10,000  g, 15 min, 25  C; swinging buckets) over precision coverslips (see Notes 2 and 13). 2. Fix samples with fixation solution for 5 min, R/T. 3. Treat coverslips with antifading solution for 10 min, R/T. 4. Block samples with blocking solution for 1 h R/T. 5. Stain for external proteins with specific primary antibodies and corresponding secondary antibodies or fluorescent adducts 1 h at R/T if primary antibodies are unconjugated. Secondary antibodies should be highly cross-absorbed and assayed for noise-to-signal ratio and background. Usually detected proteins are tetraspanins. Antibodies are diluted in 1.5% BSA in HBSS. 6. Permeabilize samples with the permeabilizing solution and treat again for autofluorescence with the antifading solution. 7. Stain intraluminal components, such as anti-DNA (0.001 μg/ ml) or anti-Tsg101 (20 μg/ml) mouse fluorochrome- or biotin-conjugated monoclonal antibodies for 1–3 h at 4  C,

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followed by Alexa-488-conjugated streptavidin. Avoid unconjugated mouse-monoclonal antibodies if already used for external proteins. 8. Extensively wash coverslips after each step with HBSS, and add a final wash with deionized water before mounting. 9. Mount samples on a water-based mounting medium. Use spacers to allow enough water-based medium for imaging under TIRF microscope. 3.5.2 Preparation of Samples on Capture Antibodies-Coated Surfaces(Fig. 1c)

1. Apply around 8  106 isolated exosomal particles over precision coverslips or glass-bottom chambers (see Note 2). 2. Allow particles to bind to antibodies for 30 min to 1 h at 4  C (see Note 14). 3. Gently wash with HBSS twice. 4. Fix samples with fixation solution for 5 min, R/T. 5. Treat coverslips with antifading solution for 10 min, R/T. 6. Block samples with blocking solution for 1 h R/T. 7. Avoid use of unconjugated mouse-monoclonal antibodies. Stain for external proteins with specific primary antibodies and corresponding secondary antibodies or fluorescent adducts 1 h at R/T if primary antibodies other than mouse monoclonal are unconjugated. Secondary antibodies should be highly cross-absorbed and assayed for noise-to-signal ratio and background. Usually detected proteins are tetraspanins. Antibodies are diluted in 1.5% BSA in HBSS. 8. Permeabilize samples with the permeabilizing solution and treat again for autofluorescence with the antifading solution. 9. Stain with anti-DNA (0.001 μg/ml) or anti-Tsg101 (20 μg/ ml) mouse monoclonal antibodies for 3 h at 4  C, followed by highly cross-absorbed Alexa-488-conjugated goat anti-mouse antibody. 10. Extensively wash coverslips after each step with HBSS, and add a final wash with deionized water before mounting. 11. Mount samples on any water-based reagents such as ProLong Gold Antifade Mountant or Citifluor Phosphate-Buffered Saline AF3 with antifading for glass-bottom chambers. Use spacers to allow enough water-based medium for imaging under TIRF microscope in the case of coverslips, mounted with ProLong Gold Antifade Mountant or homemade Mowiol-based mounting medium.

3.6 Imaging and Analysis Under TIRF Microscope

Samples are imaged under a TIRF microscope with equal experimental conditions for image acquisition for the different samples to be compared, either in coverslips or glass-bottom chambers (see Note 15).

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1. Use a penetration depth of 90 nm to avoid observation of more than one vesicle in the volume. 2. A 2 zoom should be used to obtain an optical magnification. 3. Laser intensity should be maintained constant. Allow lasers or diode to be switched on for 1 h to have stable emission. 4. Acquire images including similar number of pixels or ROI (region of interest) and exposure time (see Fig. 3a). 5. Open images with Image J or a similar software for processing and analysis. 6. The surface plot plugging allows the visualization of the overlay, including pseudocolors corresponding to the different proteins detected (see Fig. 3b). 7. Concatenate all the images in different stacks according to the fluorochromes to be analyzed. 8. Brightness and contrast adjustments should be equal for all images: normalize the complete generated stack. 9. Make an overlay stack of the images for the different channels. 10. Use the scattered plot plugging to detect the mean fluorescence intensity and area of the particles in the images. Save the data in a .txt file. 11. Generate graphs in any software allowing processing of high amount of data to observe the correspondence between signals (see Fig. 3c).

4

Notes 1. Human and mouse IL-2 are functionally interchangeable. In our hands, huIL-2 is more stable and efficient for mouse cell culture than the other way around. 2. Use 10 or 13 mm diameter coverslips to reduce the amount of antibodies or substrate used. 3. To deplete FCS of exosomes, it is pretreated as follows: ultracentrifuge at 100,000  g, 4  C, o/n. Sterile filter the supernatant by passing the content of each ultracentrifuge tube through a vacuum-connected 0.22 μm filter. Stock is stored at 20  C and used to supplement medium as normal FCS (10%), for use as exosome-production medium. 4. Human γ-globulins are used in the cell sample for immunofluorescence protocols to block any unspecific cross-reaction mediated by lateral interactions of antibodies with other molecules or with Fc receptors present. 5. A 100 objective with a high NA is required to acquire images by TIRFM with high resolution.

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Fig. 3 Exosome analysis through TIRF microscopy. (a) Images from TIRF microscopy. Detected molecules are indicated. Overlay and single color images are shown. Bar, 10 μm. (b) Graphs generated with the surface plot plugging from Image J showing correspondence of signal from detected molecules. (c) Graphs showing the correlation between mean fluorescence intensity analyses of the images

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6. Do not use antibody and magnetic beads isolation to purify specific populations at this point. The small proportion of APCs will be used to present the SEE to CD3+ positive cells (SEE in solution favors apoptosis). 7. SEE binds to the extracellular β2 loop of the MHC class II and specifically to the extracellular Vβ8 region on the TCR, acting as a clamp between the APC and the T cell to activate the T cell in a specific manner. 8. Cells will show a polarized shape, with a uropod-like trailing edge, on the fifth day from stimulation. The amount of polarized cell extent of observed cell population indicates the quantity of Vβ8+ CD3+ cells present in the cell culture and depends mostly on the donor and on the treatment, with about 20% of cells at this stage. 9. We recommend including a negative control for exosome presence without cells to analyze the medium used during cell culture and exosome production, which will be processed in parallel with the rest of the samples. 10. ImageQuant LAS-4000 chemiluminescence and fluorescence imaging system (Fujifilm). 11. If you observe a low signal-to-noise ratio during imaging of your preparations due to high background, centrifuge your immunofluorescence solutions and buffers at 22  C. 12. Alternatively, you can incubate poly-L-Lys for 1 h at 37  C, unless you observe an increased background. 13. To centrifuge the coverslips, you will need plexiglass adaptors for round-bottom tubes [12]. 14. Exosomes can be incubated o/n at 4  C if you detect low binding of particles. 15. The evanescent field is detected through TIRFM imaging very near the objective, with a working distance range approximately between 70 and 300 nm, with an optimal distance of 150 nm; 90 nm allows narrowing the imaging volume to the exosome size.

Acknowledgments This work has been funded by grant SAF2017-82886-R from the Spanish Ministry of Economy and Competitiveness (MINECO), grant S2017/BMD-3671-INFLAMUNE-CM from the Comunidad de Madrid, a grant from the Ramo´n Areces Foundation “Ciencias de la Vida y la Salud” (CIVP19A5941 XIX Concurso-2018), and a grant from Ayudas Fundacio´n BBVA a Equipos de Investigacio´n Cientı´fica ( BIOMEDICINA-2018), ‘La Caixa’ Banking

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Foundation (HR17-00016) and grants BIOIMID (PIE13/041) and CIBER Cardiovascular (CB16/11/00272) from Fondo de Investigacio´n Sanitaria del Instituto de Salud Carlos III and cofunding by Fondo Europeo de Desarrollo Regional (FEDER). The Centro Nacional de Investigaciones Cardiovasculares (CNIC) is supported by the Spanish Ministry of Economy and Competitiveness (MINECO) and the Pro-CNIC Foundation and is a Severo Ochoa Center of Excellence (MINECO award SEV-2015-0505). Authors declare no competing interest. Funding agencies have not participated in the design of the studies, with no copyright over the study. References 1. Raposo G, Stahl PD (2019) Extracellular vesicles: a new communication paradigm? Nat Rev Mol Cell Biol 20(9):509–510 2. Maas SLN, Breakfield XO, Weaver AM (2017) Extracellular vesicles: unique intercellular delivery vehicles. Trends Cell Biol 27 (3):172–188 3. Raposo G, Stoorvogel W (2013) Extracellular vesicles: exosomes, microvesicles, and friends. J Cell Biol 200(4):373–383 4. Correa E, Caballero Z, De Leon LF, Spadafora C (2020) Extracellular vesicles could carry an evolutionary footprint in interkingdom communication. Front Cell Infect Microbiol 10:76. https://doi.org/10.3389/fcimb. 2020.00076 5. Thery C, Ostrowski M, Segura E (2009) Membrane vesicles as conveyors of immune responses. Nat Rev Immunol 9:581–593 6. Mittelbrunn M, Sanchez-Madrid F (2012) Intercellular communication: diverse structures for exchange of genetic information. Nat Rev Mol Cell Biol 13:328–335 ˜ez-Mo´ M, Siljander PRM, Andreu Z et al 7. Ya´n (2015) Biological properties of extracellular vesicles and their physiological functions. J

Extracell Vesicles 4:1–60. https://doi.org/10. 3402/jev.v4.27066 8. Kalra H, Richard JA, Aikawa E et al (2012) Compendium for extracellular vesicles with continuous community annotation. PLoS Biol 10:1–5. https://doi.org/10.1371/journal. pbio.1001450 9. Villarroya-Beltri C, Gutie´rrez-Va´zquez C, Sanchez-Madrid F, Mittelbrunn M (2013) Analysis of microRNA and protein transfer by exosomes during an immune synapse. Methods Mol Biol 1024:41–51 10. Calvo V, Izquierdo M (2018) Imaging polarized secretory traffic at the immune synapse in living T lymphocytes. Front Immunol 9:684 11. Torralba D, Baixauli F, Villarroya-Beltri C et al (2018) Priming of dendritic cells by DNA-containing extracellular vesicles from activated T cells through antigen-driven contacts. Nat Commun 9(1):2658. https://doi. org/10.1038/s41467-018-05077-9 12. Boleti H, Karsenti E, Vernos I (2001) The use of dominant negative mutants to study the function of mitotic motors in the in vitro spindle assembly assay in Xenopus egg extracts. Methods Mol Biol 164:173–189

Methods in Molecular Biology (2021) 2346: 105–120 DOI 10.1007/7651_2020_321 © Springer Science+Business Media New York 2020 Published online: 09 September 2020

Rapid Visualization of Intracellular Vesicle Events During Synaptic Stimulation Noa B. Martı´n-Co´freces, Amelia Rojas-Gomez, Sara G. Dosil, Irene Fernandez-Delgado, and Francisco Sa´nchez-Madrid Abstract The immune synapse (IS) enables cell-cell communication between immune cells through close contacts, as well as T-cell activation and vesicle secretion. It is sustained by fine-tuned molecular interactions of receptors at both cell sides of the IS and intracellular cytoskeletal components. The resulting intracellular polarization of different organelles, through cytoskeleton-guided vesicular traffic, is a key player in IS formation and signaling. We describe herein a method to analyze rapid changes of vesicle localization through microscopy analysis upon polarization toward the IS. These vesicles are monitored using the centrosome and its associated microtubular network or the actin-based structures as spatial references during the organization of the IS. Keywords Immune synapse, Cytoskeleton, Signaling, T-cell receptor, Vesicles, Endosomes, Actin

1

Introduction The immune synapse (IS) is a specialized cell-to-cell contact that allows the communication between the T cell and the antigenpresenting cell (APC). This highly organized structure enables the relocation of different membrane receptors such as the T-cell receptor (TCR), integrin receptors, and the activation of associated downstream signaling pathways [1, 2]. The actin- and tubulinbased cytoskeletons undergo dramatic changes, helping the narrowing of the extracellular space into the synaptic cleft and cell internal rearrangement. The actin and microtubular networks reorganize toward the IS and regulate the polarization of several organelles around the centrosome, such as the Golgi apparatus (GA); the endolysosomal system, including multivesicular bodies (MVB); and mitochondria [3, 4]. The intense vesicular traffic observed at the IS is mainly driven by the microtubules beneath the plasma membrane, while actin-related structures mediate the

Sa´nchez-Madrid and Martı´n-Co´freces contributed equally to this work.

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reorganization of the receptors on the cell surface. Actin cytoskeleton also regulates exocytosis sites at the IS [1, 5–8]. Here, we describe several microscopy-based approaches to assess dynamic changes in the cell during the IS. These include the distribution of vesicles and vesicle-associated structures related to actin dynamics, cell adhesion, and T-cell activation in response to specific antigen stimulation. To carry out this assessment, we take advantage of different protein markers and methodologies that allow the study of TCR activation and IS formation.

2

Materials

2.1 Primary Cells and Cell Lines

1. Primary T lymphocytes from human healthy donors (purified CD4+, SEE- or PHA-specific blasts). 2. Primary CD4+T lymphocytes from mouse lymph nodes and spleen. 3. Jurkat lymphoblastoid cell lines; E1–6 (Vαl.2Vβ8+TCR) or CH7C17 cells (HA1.7Vβ3+transgenic αβTCR, specific for HA peptide).

2.2

Reagents

1. Flasks for cell culture. 2. Petri dishes for protocols of primary cell isolation including cell adhesion. 3. Cytokines: human recombinant IL-2 cytokine. Mouse or human recombinant IL-7. 4. β-mercaptoethanol for cell culture. 5. 70 μm cell strainers. 6. Monoclonal antibodies for stimulation. Human cells: purified OKT3 (eBiosciences), T3b (produced at the laboratory), or HIT-3a (Biolegend) monoclonal antibodies for CD3ε and CD28.2 for CD28 (BD Biosciences). Mouse cells: 2c11 clone for CD3ε and clone 37.51 for CD28 (BD Biosciences). Other mouse T-cell activators: 50 ng/mL Phorbol 12-myristate 13-acetate (PMA) and 500 ng/mL ionomycin; 5 μg/mL OVA323-339 peptide or 2 μg/mL of concanavalin A (ConA). 7. Recombinant proteins for stimulation: recombinant ICAM1Fc of human or mouse origin (R&D; ICAM-1-Fc from human origin is available to be produced at the laboratory). 8. Staphylococcus aureus Technologies).

enterotoxin

E

(SEE;

Toxin

9. Plasmids: speckle actin-mCherry [5], CD3ζ-GFP, CD3ζ-mCherry (kind gift from Dr. B Alarco´n) [6], EB3-GFP, and EB3-RFP (kind gift from Dr. A Akhmanova [6–8]).

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10. Highly adherent poly-L-lysine hydrobromide 75,000 > Mw > 150,000, γ-irradiated for cell culture. 11. Coverslip-bottom chambers home-made or commercial (35 mm diameter iBIDI or Mat-Tek Corporation). Use No. 1.5 or 1.5H thickness to optimize image quality (see Note 1). 2.3

Media

1. Complete medium: RPMI 1640 supplemented with FCS (fetal calf serum; 10%), glutamine (100 mM), nonessential amino acids, HEPES (25 mM). Complement medium for mouse cells with β-mercaptoethanol (1 mM). 2. Incomplete medium: RPMI 1640, HEPES (25 mM), L-glutamine (100 mM), nonessential amino acids. 3. Wash solution: Hank’s balanced salt medium (HBSS). 4. Isolation wash solution: 1% FCS, 1 mM EDTA in HBSS. 5. Saline solution: NaCl (154 mM) in deionized water. 6. Transfection Scientific).

medium:

Opti-MEM

I

(Thermo

Fisher

7. Lymphocyte separation medium: any commercial density gradient media such as Biocoll or Ficoll Histopaque. 8. Coating bicarbonate-carbonate buffer for stimulating surfaces: NaHCO3 (0.1 M), Na2CO3 (0.032 M), pH: 9.6. 9. Imaging medium: 1% FCS, 25 mM HEPES (pH: 7.4) in HBSS. 10. Fixation solution: stock of 4% paraformaldehyde (PFA) in HBSS including 0.12 M sucrose. 11. TBS (Tris-buffered saline): Tris-HCl 50 mM (pH: 7.4), NaCl (154 mM). 12. PHEM (2): 120 mM Pipes, 50 mM HEPES, 20 mM EGTA, 4 mM MgCl2; pH 6.9. 13. Lysis buffer: Tris-HCl 50 mM pH ¼ 7.4 including 150 mM NaCl, 1.5 mM MgCl2, 1% NP40, 0.2% Triton X-100, 2 mM EDTA, and protease and phosphatase inhibitors. 14. Erythrocyte lysis buffer: any home-made or commercial buffer such as ACK (Thermo Fisher). 15. T-cell isolation solution: PBS (phosphate-buffered saline), 0.5% Bovine serum albumin (BSA), and 5 mM EDTA. 2.4

Equipment

1. Amaxa Nucleofector. 2. BioRad Gene-pulse Electroporator. 3. Confocal imaging: confocal and higher-resolution 3D imaging. Lightning and Navigator packages included: TCS SP8 Navigator confocal laser scanning unit with spectral detection and

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resonant scanner, equipped with a WLL laser (range 470–670 nm), attached to an inverted epifluorescence microscope (DMI6000) fitted with an HC PL Apo CS2 63x/1.4 OIL objective (see Note 2). Microscope is mounted into microscope environmental chamber with heat (temperature regulator TempControl-37-2 digital) and humidity and CO2 gas controllers (CTI-Controller 3,700 digital). 2.5

Software

1. LAS X 3.1.1. 15751 for image acquisition and analysis. 3D viewer. 2. Imaris software 7.2.2 (Bitplane) for image analysis or more advanced editions of the software. 3. Any statistical and data processing program.

3

Methods

3.1 Isolation of T Cells 3.1.1 Isolation of Human PBLs

PBLs (peripheral blood lymphocytes) may be isolated from complete blood (50–200 mL) or Buffy coat preparations (50 mL from 450 mL of peripheral blood) from healthy human donors. Lymphoblasts are more prone to be transfected than directly isolated cells. 1. Dilute the buffy coat preparation (1:3) or the complete blood (1:1) with HBSS or saline solution, and mount the gradient over a lymphocyte separation medium cushion (1:2, v/v). 2. Spin the gradient (700  g, 30 min, R/T) w/o brake. 3. Once the cells are recovered from the interphase, wash them with saline solution or HBSS four to six times to drain the platelets by centrifugation (300  g, 10 min, R/T). 4. Deplete monocytes and granulocytes by plate adhesion in complete medium (two rounds at least) for SEE-specific or PHA lymphoblast generation (see Note 3) or purify CD3+CD4+ cells by magnetic bead-based negative selection for subsequent transfection (see Subheading 3.3). A cocktail of antibodies and streptavidin-conjugated beads is recommended (StemCell or Miltenyi Biotech).

3.1.2 Isolation of Mouse CD4+ T Cells

CD4+ T lymphocytes may be isolated from the spleen or pool of peripheral lymph nodes (mesenteric, auricular, maxilar, brachial, axillar, inguinal, and others). Mouse naı¨ve T cells are more difficult to transfect; we recommend a pre-activation step for higher transfection or nucleofection efficiency. Mouse T cells can also be transfected immediately after purification with the magnetic bead-based method without activation, with lower efficiency. Mouse T-cell activation can be performed with O/N treatment with CD3/CD28 antibodies, PMA/ionomycin, OVA323-339 peptide

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(only for T cells isolated from OT-II transgenic mice), or ConA (48-h treatment). 1. Grind lymphoid organs through a 70 μm cell strainer with T-cell isolation solution (500  g, 5 min, R/T). 2. Add 1 mL of erythrocyte lysis buffer per spleen cell suspension (3–5 min, R/T). 3. Add T-cell isolation solution or complete medium to stop cell lysis. 4. Purify CD3+CD4+ cells by magnetic bead-based negative selection for subsequent transfection (see Subheading 3.3). A cocktail of antibodies and streptavidin-conjugated beads is recommended (StemCell or Miltenyi Biotech). Purification step is recommended after OVA323-339 peptide treatment as APCs are needed for activation. 5. For longer culture time, count and grow them at 2  106 per mL in complete medium + IL-2 (200–500 Units/mL). Add IL-2 every 2 days up to 6 days. 3.2 Generation of SEE-Specific or PHA Lymphoblasts from Human PBLs

1. Count cells after recovering from adhesion step and plate them at 2  106 per mL in complete medium. 2. Add SEE (0.01 μg/mL) or PHA (1 μg/mL), and incubate for 48–72 h (see Note 4) or 24 h, respectively. 3. Wash the cells twice with HBSS (1,000  g, 5 min, R/T). 4. Count them and grow them at 2–4  106 per mL in complete medium +IL-2 (20–50 Units/mL). Add IL-2 every 2 days. 5. Upon 8 days, SEE-treated cells can be restimulated for 18–24 h (0.1 μg/mL SEE and 0.4 μg/mL PHA). Wash them twice and let grow as above. Upon 18–24 h, they can be used for transfection (see Note 5). The percentage of Vβ8+ cells found at this stage is approximately 40–60%, as measured by flow cytometry (anti-Vβ8-FITC, BD Biosciences). 6. Restimulation with SEE and PHA is possible every 15 days. Lymphoblasts may be frozen (107 cells per mL). After cryogenization procedures, an effective transfection is less efficient, but cells can be restimulated again before transfection (see Note 6).

3.3 Transfection of T Cells 3.3.1 Electroporation of Lymphoblastoid Cell Lines and Human SEE T Lymphoblasts

1. Count the cells from cell culture. Use 10–20  106 cells per sample. 2. Spin the cells (500  g, 2 min, 4  C) and discard supernatant. 3. Wash cells twice with cold HBSS (500  g, 2 min, 4 C). 4. Wash cells with cold Opti-MEM I (500  g, 2 min, 4 C). 5. Resuspend in 400 μL of Opti-MEM I with the desired DNA or RNA. A mix of DNA and RNA can be used as well. The final

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volume should be less than 410 μL for the mix. Cells in solution may be kept at 4 C until transfection (not more than 30 min). Use a 0.4 mL cuvette. 6. Electroporate cells by a pulse of 240 V, 975 mΩ (usual time: 27.5–29 ms in a GenePulser II (Bio-Rad)). 7. Recover the cells from the cuvette, and plate them immediately in incomplete medium for 4–6 h at 2  106 cells per mL. Do not leave the samples for a long time in the cuvette after electroporation. To avoid cell death, up to ten samples may be electroporated at once. 8. Add FCS at 5% for the first 18–24 h and then supplemented to 10%. 9. Supplement with IL-2 primary T lymphoblasts (10–50 U/mL). 3.3.2 Nucleofection of Human T Lymphoblasts

1. Pre-warm the nucleofection solution, the cuvettes, the cell culture medium post-transfection, and cell culture plates before nucleofection. 2. Count the cells. Use a range of 106–107 cells per nucleofection. 3. Spin the cells and discard supernatant (500  g, 2 min, 4 C). 4. Wash cells in cold HBSS twice (500  g, 2 min, 4 C). 5. Aspirate all the washing media. Add the pre-warmed transfection mix directly over the cells (VPA-1002 including the DNA or RNA) (see Note 6). 6. Put the cell mix into the pre-warmed cuvette to be nucleofected immediately. Do not leave the cells with the transfection medium for more than 3–5 min. 7. Use program T23 (Nucleofector I-Amaxa, see Note 7). 8. Nucleofect a single point, and plate cells immediately after the pulse in incomplete medium for 4–6 h at 2  106 cells per mL. Then supplement with FCS (10%) and IL-2 (20 U/mL).

3.3.3 Nucleofection of Mouse CD4+ T Cells

1. Pre-warm the nucleofection solution, the cuvettes, the cell culture medium post-transfection, and cell culture plates before nucleofection. 2. Count the cells. Use 5  106 cells maximum per nucleofection. 3. Spin the cells and discard supernatant (500  g, 2 min, 4  C). 4. Wash cells twice in cold HBSS (500  g, 2 min, 4  C). 5. Aspirate all the washing media, and add 100 μL of the pre-warmed transfection mix: Opti-MEM I with the DNA or RNA included (see Note 6). 6. Add the cell mix to the pre-warmed cuvette and nucleofect immediately. 7. Use program X-01 (Nucleofector I-Amaxa, see Note 7).

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8. Nucleofect a single cuvette, recover the cells, and plate them immediately in complete medium at 4  106 cells per mL supplemented with recombinant mouse IL-7 (5 ng/mL). Do not keep cells in the nucleofection solution for more than 3–5 min. 3.4 Coating of Coverslip-Bottom Chambers

1. Pipette 50–100 μL of poly-L-lysine hydrobromide in sterile water (50 μg/mL) per chamber and incubate o/n at 4  C.

3.4.1 Preparation of poly-L-Lys-Coated Surfaces

2. Wash twice with sterile deionized water.

3.4.2 Preparation of Stimulating Surfaces

1. Mix the corresponding anti-CD3 and anti-CD28 (3:1 ratio) antibodies in coating buffer. Add ICAM1-Fc (1 μg/mL). These antibodies and recombinant proteins do not cross-react.

3. Store in imaging medium until used. They can be frozen at this step. Do not allow to dry.

2. Pipette 50–100 μL of the mix per dish and incubate o/n at 4  C. Alternatively, 1–2 h at 37 C may be used, but the first method is preferred. 3. Wash twice with pre-warmed HBSS. 4. Pre-warm at 37 C in imaging medium before use. Plates can be frozen at this step. Do not allow to dry. 3.5 Live Imaging Vesicular Traffic Studies

1. Transfect T cells with speckle actin-mCherry or EB3-RFP and CD3ζ-GFP (or EB3-GFP combined with EB3-RFP) plasmids 24 h before imaging.

3.5.1 Preparation of T Cells

2. Collect transfected cells, spin them (500  g, 5 min, R/T), and resuspend in 2 mL of HBSS. 3. To recover live cells, add 1 mL of lymphocyte separation medium cushion to the bottom of the tube. Spin cells without brake (1200  g, 5 min, R/T). Live cells relocate at the interface with the lymphocyte separation medium, whereas dead cells accumulate at the bottom of the tube. 4. Cells can be sorted at this point to allow homogeneous expression of the fluorescent proteins. Spin the cells after sorting, and keep them in the cell incubator with complete medium 2 h before imaging. 5. Count the cells. Use 5  105 cells for each video. 6. After each knock-down has been performed, keep 5  105 cells per well of a mini-gel for immunoblot confirmation, and use lysis buffer (40 μL per 106 cells) to process samples through native or SDS-PAGE and transfer to nitrocellulose or PVDF membrane.

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7. Wash cells twice with HBSS, and resuspend in imaging medium (107 cells per ml). Pre-warm at 37  C and 5% CO2 until used (see Note 8). 8. Pre-warm control and stimulating dishes with 2 mL of imaging medium before use. 3.5.2 Image Acquisition

1. Pre-warm at 37 C, and adjust CO2 to 5% and humidity of the confocal stage for at least 1–2 h before image acquisition. Also, pre-warm immersion media. Put a drop of it onto the objective (63; 1.4 NA; maximal zoom to be used is 3 in the Leica microscope). Introduce the pre-warmed dish in the stage. Add 10 μL of cells, and localize the transfected cells with the oculars. Place cells at the center of the imaging area. 2. To allow high-speed scanning, use the resonant scanner (8000 Hz) and the bidirectional scanning mode. Use 512  512 px size to reduce acquisition time. Use the mode of scanning “between lines” for the acquisition of different fluorophores, if your fluorochromes can be detected. 3. Acquire CD3ζ-GFP, speckle actin-mCherry, or EB3-RFP fluorescence and bright field at same time (see Note 9). 4. It will require accumulating signal and using a low-intensity of laser to avoid cell toxicity. 5. Include the whole cell in the Z stack (about 10–15 μm). Take images every 0.25–0.5 μm. This will allow a framelapse of about 1–1.2 s. 6. Cells are rapidly adhered to the surface, allowing the formation of an actin-based lamella during a spreading that increases its area for 1–2 min. After this event, the lamella is stable for a time, and the vesicles are relocated to the immune synapselike area. 7. Acquire time-lapse for 3–5 min. The images that can be obtained are observed in Fig. 1 and Table 1.

3.5.3 Image Analysis LAS X Software (See Fig. 1)

1. Open the file from the confocal microscope with LAS X software. Crop time and XY dimensions from the fluorescence time-lapse to analyze only the desired time range and cell of interest (or part of a cell). Probably, a unique cell has been imaged per file. 2. Image visualization may require adjustments such as background subtraction and Gaussian or median filtering for better detection of the objects from confocal Z stack time-lapse (see Note 10).

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Fig. 1 Resonant scanner imaging of vesicle and cytoskeleton dynamics. (a) Jurkat cells were co-transfected with EB3-GFP (green) and CD3ζ-mCherry (magenta) and imaged during IS formation. A maximal projection of the Z stack is shown. Note that vesicles are near the tubulin structures. (b) Jurkat cells were co-transfected with EB3-GFP and CD3ζ-mCherry and imaged during IS formation. Two different frames of a 3D reconstruction including a whole cell from the time-lapse performed with Leica accompanying software are shown. EB3-decorated tips are observed. Color legend indicates the distance from top (blue) to bottom (red) of the cell. Bottom corresponds to IS. Each stack was acquired every 0.890 s. White arrow, centrosome. (c) Jurkat cells were co-transfected with speckle actin-mCherry and CD3ζ-GFP (magenta) and imaged during IS formation. Two different orientations of same 3D reconstruction performed with Leica accompanying software are shown. Two different frames of the time lapse are shown. Color legend as in (b) Bar, 5 μm

3. A maximal projection can be obtained to have a global idea of the relative localization of intracellular structures (see Fig. 1a). Pseudocolors can be applied (see Note 11). 4. Single stacks or maximal projections of a group of stacks can help to analyze different parts of the cells, such as the bottom (structures related to cell adhesion or to synapse formation, depending on the substrate used). 5. Open the 3D viewer. The range of each channel can be adjusted. Use the “volume render” option to obtain a reconstruction of structures (maximum and shadow projections are also available). The “depth coding” scale allows easy

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Table 1 Parameters of image acquisition and analysis of vesicle dynamics. Imaris software Image acquisition Number of frames

7560

Number of channels

3

Number of slices per time

24

Frame interval

0.94963 s

Width

74.0344 μm

X resolution: 6.9157 pixels per μm

Height

43.6687 μm

Y resolution: 6.9157 pixels per μm

Depth

12.9204 μm

Voxel size: 0.1446  0.1446  0.5383 μm3

Calculated parameters Number of tracks

368

Number of spots (vesicles)

11,890

Parameter

Mean  SD

Median

Mean radius

0.760  0.4177 μm

0.750 μm

Displacement

1.952  0.4177 μm

0.911 μm

Duration

30.427  33.147 s

Speed

0.122  0.192 μm s

13.300 s 1

0.056 μm s1

visualization of general localization from bottom to top of structures (see Fig. 1b, c). 6. Orientation of the cell can be defined by the user. Visualization of single channels and merge options will allow evaluation of the complete scenario. 7. Single images in different formats can be generated (Tiff, jpg, png), as well as time-lapse movies with different options, such as spatial rotation and kinetics (avi, mpg, wmm; see Note 12). 3.5.4 Image Analysis Imaris Bitplane Software (See Fig. 2)

1. Open the file from the confocal microscope with Imaris software. Crop time and XY dimensions from the fluorescence time-lapse to analyze only the desired time range and cell of interest. Probably, a unique cell has been recorded. Usually, 10–20 s tracking is enough to have reproducible results. 2. Select the “Image processing” menu and the “Thresholding” options to perform a background subtraction and “Image processing” and the “Smoothing” options for Gaussian filtering. This will allow better detection of the objects from confocal Z stack time-lapse (see Note 10).

Fig. 2 Analysis of vesicle dynamics in whole cells. (a) Whole cell volume (cyan) generated from detected actin fluorescence (green) through the Imaris surface tools for transfected Jurkat cells. Tracks represent vesicle detection through the spots tool. Color scale for time is shown (0–17 s). Different angles are shown (b). Partial whole cell reconstruction as in (a), allowing observation of tracks and vesicles enriched in CD3ζ (red fluorescence and spots). (c) Basal tracks, corresponding to the bottom part of the cell as in (a), showing vesicle movement at the cell adhesion structure, in this case, the immune synapse. (d) Same view as in (c), but including green fluorescence from actin. (e) The same cell is analyzed as in Fig. 1c with the LAS X software, for comparison. Color scale corresponds to distance to top of the cell (blue); bottom of the cell corresponds to immune synapse (red)

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3. To analyze the dynamics of the vesicles, use the “Spots” utility. Select the manual adjustment of parameters and the “Track surfaces over time” option. 4. Select the corresponding channel for signal detection, and allow region growing option. Estimated XY diameter can be measured with the “Slice” view, and then return to the “surpass” view. If you observe that the vesicles are not completely spherical, select “ellipsoid detection” and estimate the Z diameter. 5. Enable “Background subtraction,” and use a “Quality” parameter allowing specific detection of the vesicles (the value will depend on your signal-to-noise ratio). The region-growing parameter rendering better results to identify objects is “Local contrast” option. 6. Select the “Autoregressive Motion” for tracking algorithm, selecting a maximal distance of about 0.5–1 μm. Allow a maximal gap size of 0 or 1. 7. Select a “Track duration” above three frames (the time corresponds to the interval between frames). 8. The generated tracks can be manually revised. Vesicles can be visualized in different color scales as spots for their calculated volume. The tracks are shown in a time scale. 9. Export and save the spot creation parameters for bulk analysis of data. 10. For statistics, export in any data file format, such as excel; number, localization, length, displacement length, and duration parameters from the tracks, among others, are provided. See Table 1. 11. For analysis of cytoskeleton dynamics, create a new channel for speckle actin-mCherry, EB3-GFP, or EB3-RFP tracking with the “Surface” tool if you aim to observe the volumes corresponding to cytoskeletal components (see Fig. 2 and [9] for EB3-tips). 12. Select the manual adjustment of parameters and the “Track surfaces over time” option for microtubule tips. Tracking actin surfaces is not recommended. Select the “Smooth” option to measure the dimensions of the object identified. The large diameter parameter indicates the larger size of the object that fits into the surfaces to be detected through the “Background subtraction” option. Select “Surface area detail” of one-tenth of the maximal diameter as a regular measure. 13. Adjust the threshold to allow detection of individualized surfaces. Indicate a number of voxels to determine the size of the particles included in the analysis (usually about 3–5). Select the “autoregressive motion” algorithm to calculate the tracks for

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detected surfaces. Indicate the maximal distance between surfaces in a track (usually 0.5–0.75 μm) and the maximal gap to consider a unique track (0 or 1). The minimal duration of a track to be included in the analysis is usually 3–5 frames (about 1.5–3 s). 14. Tracks can be manually adjusted. 15. For calculation of cell surface, select the “Surface” tool; do not track it. At the “Smooth” option, the value for the surface area detail is usually the same as the one used before for tubulin- or actin-generated surface, but the major cell diameter is to be used as the value for the maximal size of the object that fits into the surfaces to be detected in the “Background subtraction” option (10–20 μm depending on the cell size). The minimal number of voxels should be 30 or more to avoid little surfaces. Delete manually those surfaces that do not represent the cell volume. 16. Export and save the surface creation parameters for bulk analysis of data. 17. For statistics, export in any data file format, such as excel; number, localization, length, displacement length, and duration parameters from the tracks, among others, are provided (not recommended for actin). 18. For statistics, use the sum of intensities from cytoskeletongenerated surfaces and from whole cell-generated surfaces to calculate the ratio of incorporated fluorescence into actin or tubulin structures per frame as a measure for actin or tubulin polymerization, respectively (see Note 10).

4

Notes 1. Use 10 or 13 mm diameter coverslips to optimize and reduce the amount of antibodies or substrate used. 2. Confocal imaging with a 63 objective with a high NA is recommended for better resolution. The use of a 100 objective will improve the resolution of the image, but will absolutely need a stronger signal from fluorochromes to be detected. Therefore, using the optical zoom and the crop option available from your system can be a better option to increase time resolution and signal-to-noise ratio. 3. Do not use biotin-conjugated antibodies and streptavidinconjugated magnetic beads to purify relevant cells. The small proportion of APCs will present SEE to CD3+ positive cells instead of having SEE in solution, which favors apoptosis.

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4. SEE acts as a clamp between the APC and the T cell through its binding to the β2- extracellular loop of the MHC class II and to the Vβ8 extracellular region of the TCR (with more affinity than to other Vβ domains), producing a polyclonal stimulation. 5. Cells may be nucleofected every time with the Amaxa system (human T-cell kit, program T23) or electroporated in Optimem I medium (Thermo Fisher Scientific) with the Genepulser II from Bio-Rad (240 V, 975 mω), after the first restimulation with either DNA or RNA. CD3+CD4+ T cells (most of them Vβ8+) can be isolated by magnetic bead-based negative selection and transfected afterward. 6. Frequently, the transfection procedure results in about 50% cell viability. Viable cells will divide properly unless the normal cell cycle is affected by overexpressed or exogenously expressed molecules. Avoid adding more than 5 μL of DNA or RNA to 100 μL of mix. RNA is usually used at 1–4 μM. Optimized range: 1.5–2.5 μL in 100 μL of Amaxa mix. 7. Nucleofect and plate all cells at once in a single mix with warmed medium in a cell culture dish or flask. This will improve cell survival. 8. Cells can be sorted before imaging, if the transfection efficiency is very low, to improve timing and microscope use. 9. Hybrid detectors are a good idea to increase signal-to-noise detection; however, they can detect very low intensity, depending on the expression of your fluorescent protein (or fluorochrome-conjugated antibody or fluorescent probe, if used in in vivo assays). Regular photomultipliers will allow increasing signal detection with posterior processing of the images. 10. Although mean fluorescence intensity (MFI) measures will be rendered by the software, resonant scanning does not allow calculating them correctly. In this case, fluorescence cell ratios can be established. A ratiometric analysis of the incorporated fluorescence in actin- or tubulin-related structures vs the whole cell indicates the amount of actin or tubulin incorporated in structures (i.e., polymerization). Increases or decreases in these ratios along time indicate the speed of polymerization. 11. Most scientific journals will ask to compose figures in colors that allow observation by color blind people. A green and magenta combination is common; providing single channels will also facilitate visualization. Black and white scales form the original signal rendered by detectors. 12. The program uses different codes for movie formats; use the one that allows visualization in your media. “Avi raw” format is recommended to record all the information on the images,

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although it generates heavy files, since it can be easily used in other software such as Image J (National Institutes of Health: http://rsb.info.nih.gov/ij/).

Acknowledgments We want to thank the dedication of the staff of the Microscopy Facilities and their effort to help during the special situation created by the pandemic outbreak. Optical microscopy experimentation has been conducted at the Microscopy and Dynamic Imaging Unit of the CNIC (Centro Nacional de Investigaciones Cardiovasculares) and at the Microscopy Facility of the IIS-IP (Instituto Investigacio´n Sanitaria-Instituto Princesa), Madrid, Spain. This work has been funded by grant SAF2017-82886-R from the Spanish Ministry of Economy and Competitiveness (MINECO), grant S2017/BMD3671-INFLAMUNE-CM from the Comunidad de Madrid, a grant from the Ramo´n Areces Foundation “Ciencias de la Vida y la Salud” (CIVP19A5941 XIX Concurso-2018), and a grant from Ayudas Fundacio´n BBVA a Equipos de Investigacio´n Cientı´fica ( BIOMEDI CINA-2018), and “La Caixa” Banking Foundation (HR17-00016). BIOIMID (PIE13/041) from Instituto de Salud Carlos III, CIBER Cardiovascular (CB16/11/00272, Fondo de Investigacio´n Sanitaria del Instituto de Salud Carlos III and co-funding by Fondo Europeo de Desarrollo Regional FEDER). The Centro Nacional de Investigaciones Cardiovasculares (CNIC) is supported by the Spanish Ministry of Economy and Competitiveness (MINECO) and the Pro-CNIC Foundation and is a Severo Ochoa Center of Excellence (MINECO award SEV-2015-0505). The authors declare no competing interest. Funding agencies have not participated in the design of the studies, with no copyright over the study. References 1. Dustin ML, Olszowy MW, Holdorf AD, Li J, Bromley S, Desai N, Widder P, Rosenberger F, van der Merwe PA, Allen PM, Shaw AS (1998) A novel adaptor protein orchestrates receptor patterning and cytoskeletal polarity in T-cell contacts. Cell 94:667–677 2. Monks CR, Freiberg BA, Kupfer H, Sciaky N, Kupfer A (1998) Three-dimensional segregation of supramolecular activation clusters in T cells. Nature 395:82–86 3. Martin-Cofreces NB, Baixauli F, SanchezMadrid F (2014) Immune synapse: conductor of orchestrated organelle movement. Trends Cell Biol 24:61–72 4. Finetti F, Patrussi L, Masi G, Onnis A, Galgano D, Lucherini OM, Pazour GJ, Baldari

CT (2014) Specific recycling receptors are targeted to the immune synapse by the intraflagellar transport system. J Cell Sci 127:1924–1937 5. Garcia-Ortiz A, Martin-Cofreces NB, Ibiza S, Ortega A, Izquierdo-Alvarez A, Trullo A, Victor VM, Calvo E, Sot B, Martinez-Ruiz A, Vazquez J, Sanchez-Madrid F, Serrador JM (2017) eNOS S-nitrosylates beta-actin on Cys374 and regulates PKC-theta at the immune synapse by impairing actin binding to profilin-1. PLoS Biol 15:e2000653 6. Martin-Cofreces NB, Baixauli F, Lopez MJ, Gil D, Monjas A, Alarcon B, Sanchez-Madrid F (2012) End-binding protein 1 controls signal propagation from the T cell receptor. EMBO J 31:4140–4152

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7. Grigoriev I, Akhmanova A (2010) Microtubule dynamics at the cell cortex probed by TIRF microscopy. Methods Cell Biol 97:91–109 8. Blas-Rus N, Bustos-Moran E, Perez de Castro I, de Carcer G, Borroto A, Camafeita E, Jorge I, Vazquez J, Alarcon B, Malumbres M, MartinCofreces NB, Sanchez-Madrid F (2016) Aurora

A drives early signalling and vesicle dynamics during T-cell activation. Nat Commun 7:11389 9. Blas-Rus N, Bustos-Moran E, Sanchez-MadridF, Martin-Cofreces NB (2017) Analysis of microtubules and microtubule-organizing center at the immune synapse. Methods Mol Biol 1584:31–50

Methods in Molecular Biology (2021) 2346: 121–134 DOI 10.1007/7651_2020_339 © Springer Science+Business Media New York 2020 Published online: 15 November 2020

Monitoring of Active Notch Signaling in Mouse Bladder Urothelium Panagiotis Karakaidos and Theodoros Rampias Abstract Notch signaling plays a crucial role in differentiation and homeostasis in a wide variety of epithelia. The tumor suppressor role of Notch in bladder urothelium is well accepted as the inactivation of this pathway due to damaging mutations in its components is associated with neoplastic transformation. Monitoring Notch signaling is therefore critical to understand how the deregulation of cell–cell communication can lead to differentiation loss and carcinogenesis. In this chapter, we provide a method to visualize active Notch signaling by the detection of the nuclear levels of Notch intracellular domain in mouse urothelium. The technique outlined below is characterized by high sensitivity and specificity and has been successfully applied to human tumor specimens. In this context, this technique could be used to characterize the molecular profile of Notch-deficient tumors and analyze the clonal expansion dynamics and the heterogeneity patterns of Notch inactivation. Key words Cancer, Immunofluorescence, Notch intracellular domain (NICD), Notch signaling, Tyramide signal amplification (TSA), Urothelium

1

Introduction One of the most, if not the best, studied cell–cell communication signaling pathways is the canonical Notch signaling pathway. Notch is a highly conserved pathway across many species and it plays major roles in a diverge set of key cellular functions including cell proliferation, death, and differentiation with profound effects in metazoan development and disease. The uniqueness of Notch, as compared to other major intercellular signaling pathways, relies on features like the highly topological constraints (activation requires direct cell-to-cell contact), dose-dependent response (lack of signal amplification means), pleiotropic outcome (developmental stage and/or cell context dependent response), and its signal transduction mode of regulated proteolysis (Notch receptors are proteolytically cleaved upon ligand activation and a fragment, the Notch intracellular domain (NICD), is imported to the nucleus) [1]. In mammals, there are four Notch receptors (Notch1–4) all of which are single-pass type I transmembrane proteins that transduce

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signals by binding to membrane bound ligands (Jagged1–2, Deltalike 1, 3, and 4) on adjacent cells. Their extracellular domain contains several tandem EGF-like repeats (29–36 in humans), a negative regulatory region (NRR), a single transmembrane region (TM), and the NICD (Fig. 1a). In canonical Notch pathway, upon ligand binding, Notch receptors undergo a conformational change enabling access of a membrane-bound ADAM metalloprotease that cleaves receptors in the juxtamembrane extracellular domain proximal to the transmembrane region) [2]. ADAM-dependent cleavage, also termed S2 cleavage is followed by a cleavage in the helical transmembrane region (S3 cleavage site after valine residue 1754 in human Notch1 receptor and after Valine 1744 in mouse receptor) that is executed by the aspartyl protease presenilin (PS) which is a central component of γ-secretase complex at the final steps of Notch activation releasing the Notch Intracellular Domain (NICD) [3] (Fig. 1b). NICD translocates (harbors two nuclear localization signals) to the nucleus where it binds to the DNA-bound protein CSL together with coactivator protein Mastermind (MAM) [4], leading to target gene activation [5]. This two-step activation mechanism by proteases is central to canonical Notch signaling. It is believed that in the absence of ligand, mature Notch receptors are held into an inactive state because the Negative Regulatory Region (NRR) inhibits Notch activation [6]. The S2 cleavage appears to be necessary for Notch activation in all organisms while the presenilin activity and the S3 cleavage seems to be quite essential for the fully Notch activation. Interestingly, the TM region that harbor the S3 cleavage site along with the onset of NICD are particularly conserved among species (Fig. 1c) and mutagenesis experiments have demonstrated that Val1744 is critical for Notch activity. Several excellent reviews cover in detail the pathway components and the activation mechanism of Notch pathway [1, 6, 7]. Nuclear presence of NICD is a marker of Notch signaling activation. However, several commercially available antibodies fail to detect NICD via immunohistochemical techniques in tissues and most of them work well only on cells (immunocytochemistry). The need, and interest, for in situ validation of Notch activity has been a challenge for years and affected the progress in several fields of mammalian developmental or cancer studies that had to turn in genetically labeled animal models. The proposed protocol offers a tool to study Notch in paraffin embedded tissue sections. In particular, we have optimized the conditions for the identification of nuclear NICD in epithelial tissues of the urinary tract. In both bladder and ureter, the lining epithelium is the same and it is known as urothelium. It consists of three cell layers, top to bottom, umbrella, intermediate and basal cells. We have previously shown, using this protocol, that Notch is active in all three cell types in both bladder and ureter epithelium [8] (Fig. 2, upper panel). As a proof

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Fig. 1 Proteolytic sites and activation of Notch signaling. (a) Schematic representation of the human Notch1 receptor structure indicating the location of S2 and S3 cleavage sites. (b) Sequence and tertiary structure of Notch1 transmembrane and associated juxtamembrane segment indicating the position of Valine 1754. WebGL-based NGL viewer was used to display the NMR structure of 5KZO RCSB PDB in three dimensions. (c) Alignment of protein sequences corresponding to the transmembrane and associated juxtamembrane domain of Notch1 receptor (segment 1720–1770 aa of human Notch1 receptor) from different species. Identical amino acids are represented as dots

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Fig. 2 Detection of nuclear NICD levels in urothelium. Immunofluorescence (IF) showing (from left to right) staining for NICD, Pankeratin, and DAPI in (Upper panel) normal bladder and ureter epithelium. (Lower panel) Ureter tumors developed in R26rtTA; tetO-Cre; Ncstnflox/flox mice that lack active Notch signaling. All scale bars, 50 μm

for the efficiency and validity of the proposed method we utilized a genetic mouse model that lack intracellular Notch (R26rtTA; tetOCre; Ncstnflox/flox), as a negative control in urothelial tissue, and we verified the specificity of the primary antibody of our protocol in developed ureter tumors (Fig. 2, lower panel). The benefits of using this protocol for immunohistochemical detection of NICD, apart from the high efficiency and specificity of nuclear Notch detection, include the reduced amount of primary antibody required, the utilization of masking antibody for sufficient histological information regarding specific topological tissue information (e.g., discrimination of epithelial and connective tissue), as well as the increased sensitivity. The latter is obtained through the use of tyramide signal amplification (TSA) which has been proven that can be applied in any primary antibody and enhance its signal-to-noise outcome many-fold, particularly for “difficult” antibodies (i.e., high background antibodies or low-copy number epitopes) [9]. TSA applications are versatile and commercially available. Of note, the primary antibody we suggest herein is raised in rabbit,

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immunized with a synthetic peptide corresponding to the conserved sequence of human Notch1 receptor that we show in Fig. 1c (1720–1770 aa) and includes the Val1754 cleavage site (equivalent to Val1744 in mouse Notch1). This antibody does not recognize the full-length Notch1 receptor and is applicable at least to human, mouse, and rat specimens. Notch is particularly active in several epithelial tissues during development and adulthood. Apart from healthy tissues, deregulated Notch signaling is implicated in several human diseases including cancer. Notch has been described as tumor suppressor or oncogene depending on the host tissue [10]. Our protocol is applicable also to cancer histological specimens for screening Notch status. To this end, the efficiency of our protocol in bladder cancer has been published [8], while herein we provide some unpublished results from a tissue microarray (YTMA 211) of human head and neck tumors (Fig. 3), in which Notch is significantly compromised due to mutations on pathway components with profound effect on Notch receptor maturation and nuclear localization [11]. Again, the costaining with the same masking antibody provides a clear distinction of the tumor area (which is positive for cytokeratins) and adjacent normal connective tissue (negative for cytokeratins) which is difficult to discriminate with DAPI counterstaining only.

2 2.1

Materials Mice

2.2 Tissue Fixation Reagents and Components

We used 10–12- weeks old wild-type C57BL/6 mice, to monitor the activity of Notch signaling in the urothelium. 37% formaldehyde (Fisher scientific, S25329). (Gas formaldehyde dissolved in water with saturation at 37–40%. Also known as 100% formalin). Use always in hood. It has been upgraded in risk to a Category 2 carcinogen. 10% neutral buffered formalin (3.7% formaldehyde, 46 mM Na2HPO4, 29 mMNaH2PO4·H2O. pH 7.0). 10% formalin actually represents 10% of the 37–40% formaldehyde stock solution. The actual amount of dissolved formaldehyde in the 10% formalin is therefore only 3.7–4.0%. Petri dish, 5.5 in. diameter (Fisher scientific, #08-747 F).

2.3 Paraffin Embedding Reagents and Equipment

Absolute ethanol (Fisher scientific, BP2818100). Xylene (Fisher scientific, X3P-1GAL). Paraffin wax (Fisher scientific, AC416770020). Microtome (Leica RM 2265). Siliconized charged glass Slides (VWR, 89500-498).

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Fig. 3 Detection of nuclear NICD levels in human cancer specimens. Immunofluorescence (IF) showing staining for NICD, Pankeratin, and DAPI in head and neck squamous cell carcinomas from tissue microarray YTMA 211. All scale bars, 50 μm

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Fig. 3 (continued) 2.4 Immunostaining Reagents

Xylene, (Laboratory Grade, Fisher Chemical). Ethanol series: 100, 95, 85, 70, 50%. Prepared from absolute ethanol. Peroxidase blocking buffer: 0.3% hydrogen peroxidase (H2O2) in methanol. Prepare fresh from 30% hydrogen peroxide solution (Fisher scientific, ACS grade, H325-100). Phosphate buffered saline, 10 solution (10 PBS): 77 mM Na2HPO4 (Sigma), 23 mM NaH2PO4 (Sigma), and 1.5 M NaCl (Sigma) in distilled water, pH 7.2. 1 PBS-T (diluent and washing buffer): 1 PBS with 0.05% (v/v) Tween 20 (Sigma). Prepare 1 L of 1 PBS and add 1 mL of Triton X-100. Used as diluent and washing buffers in conventional immunostaining. Store at 4  C.

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Antigen retrieval buffer: EDTA antigen retrieval buffer (1 mM EDTA, 0.05% Tween 20, pH 8.0). Primary and secondary antibody buffer: 0.3% (w/v) albumin bovine serum, fraction V (Sigma, A7906) in 1 PBS-T. Blocking solution: 2% BSA and 10% normal goat serum (Vector lab, S-1000-20) in1 PBS-T. Primary target antibody: Cleaved Notch1 (Val1744) (D3B8) Rabbit, (Cell Signaling, #4147). This antibody displays reactivity for human, mouse and rat species. Primary mask antibody: Mouse monoclonal [AE1/AE3] to pan-cytokeratin (Abcam, ab80826). This antibody displays reactivity for cytokeratins of mouse, rat, rabbit, chicken, cow, and human species. Secondary target antibody: HRP-labeled anti-rabbit secondary antibody: EnVision™+ System-HRP labeled polymer anti-rabbit (DAKO, K4002). Goat anti-rabbit, secondary to rabbit antiNICD antibody. Secondary mask antibody: Goat anti-mouse IgG (H+L) Alexa 488 (secondary to mouse pan-cytokeratin); (Invitrogen, A32723). Washing buffer: 1 PBS-T with 0.05% (v/v) Tween 20 (Sigma, P9416). Target fluorophore: Cy3-Tyramide reagent with amplification diluent (PerkinElmer, Boston MA, NEL744001KT). Labeled tyramide is provided as a powder and should be dissolved in HPLC grade DMSO prior to use following the kit instructions (150 μL/tube). To prepare Cy3 tyramide stock solution, dissolve the powder in DMSO inverting the vial several times. To minimize freeze–thaw cycles, stock the solution in small aliquots of 5–10 μL, depending on the quantities required for individual experiments. Store aliquots at20  C, protected from light. Mounting medium with nuclear stain: Prolong Gold antifade reagent with DAPI (Invitrogen, P36931). 2.5 Other Equipment for Immunostaining

6-quart aluminum pressure cooker (Presto). Glass Coplin staining jars. Dark slide chamber with moistened filter paper for humid incubation of slides. Cover glass, thickness 1½, 22  50 mm, #2980-225, Corning.

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2.6 Microscopy and Imaging Set Up

Upright fluorescence microscope (Leica DMRA2) with filter sets for DAPI, GFP, Cy3, Cy5, and Cy7 (Zeiss, Germany). Orca Flash 4.0 V3 CMOS Camera (Hamamatsu Photonics, Japan).

2.7 Image Processing and Analysis

Open source image processing package ImageJ (https://imagej. net/Fiji).

3

Methods We describe here the protocol for the immunofluorescent detection of NICD on paraffin embedded sections of mouse urothelium, which can be extended to any human, mouse or rat tissue. Enzyme-linked signal amplification is a key technique used to enhance the immunohistochemical detection. Tyramide signal amplification (TSA) is based on HRP catalytic deposition of labeled tyramide molecules in close vicinity to the epitope of interest. The advantages of this technique are its simplicity, enhanced sensitivity, high specificity, increased signal-to-noise ratio and compatibility with modern multilabel fluorescent microscopy [12]. Here, we describe the use of TSA technology to increase the signal of nuclear NICD on mouse urothelial tissue sections along with the method of obtaining them. Cleaved Notch1 (Val1744) polyclonal antibody detects endogenous levels of the Notch1 intracellular domain (NICD) only when released by cleavage between Gly1753 and Val1754 (equivalent to Gly1743/Val1744 of murine Notch1). This monoclonal antibody is produced by immunizing rabbits with a synthetic peptide corresponding to the sequence at the Val1754 cleavage site in human Notch1 (equivalent to Val1744 in mouse Notch1) does not recognize full-length Notch1 or Notch1 cleaved at other positions.

3.1

Tissue fixation

1. The mice are euthanized according to institutional guidelines (see Note 1). Hair on the belly is removed with an electric shaver. The lower abdomen wall is surgically opened by a midline incision up to the pubic symphysis. 2. The bladder is located in the lower center of the abdomen close to the pelvic bone. Cut the median umbilical ligament connecting the ventral surface of the bladder to the abdominal wall. 3. Hold the dome with forceps to gently pull the bladder out of the abdomen in order to separate it from other structures such as the seminal vesicles. 4. Cut the ureters as close as to the bladder and clean the bladder from fat and connective tissue.

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5. Remove the bladder from abdominal cavity by cutting the urethra. 6. Place the excised bladder immediately in in a petri dish containing cold 1 PBS (4  C). Soak it for 5 min. 7. After soaked in 1 PBS solution, the bladder is placed onto a paper towel and use a sharp straight blade to make a short cut at its equatorial line. Press the dome gently to empty the stored urine (see Note 2). 8. Place the bladder at 4  C 1 PBS solution for preparation for the next step. 9. Fix a medium size mouse bladder in 10% neutral buffered formalin, pH 7.0 (see Note 3), for 24 h (see Note 4) in 4  C (see Note 5). Ensure that the bladder is fully immersed. 10. Remove 10% neutral buffered formalin, add 70% ethanol. Store at 4  C prior to paraffin embedding. 3.2 Paraffin Embedding and Sectioning Onto Slides

Briefly, after fixation, the tissues are gently dehydrated in increasing grades of ethanol to minimize cell damage, processed through xylene to clear the tissues from ethanol, embedded in heated paraffin at 60  C, and then hardened overnight to form tissue blocks. Finally, the tissues are sectioned at a thickness of 5 μm, placed on slides and dried at room temperature. The sectioned tissues should be stored at 4  C to prevent tissue oxidation.

3.3 Immunostaining on Paraffin Sections for Detection of Nuclear Notch ICD Levels

Before proceeding with the staining protocol, tissue sections from fixed and paraffin-embedded material must first be deparaffinized and rehydrated. Incomplete removal of paraffin can cause poor staining to the section with high levels of autofluorescence.

3.3.1 Deparaffinization and Rehydration of Tissue

1. Melt the paraffin sections in a hybridization oven at 60  C for 10 min. 2. Remove residual paraffin by immersing the slides in 100% xylene for 20 min. Repeat this step one more time, gently shaking off excess liquid in between steps (see Note 6). 3. Rehydrate the slides by sequential incubation in an alcohol series, 3 min each step: 100% ethanol, 100% ethanol, 95% ethanol, 85% ethanol, 70% ethanol, 50% ethanol. It is important to not let sections dry during solution changes (see Note 7). 4. Transfer slides to tap water with a slow gradient under running water, and finally transfer the slides to a Coplin jar containing distilled water for at least 5 min for rehydration.

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1. Antigen unmasking is achieved using the heat-induced epitope retrieval (HIER) method: Fill the jar to the top with EDTA antigen retrieval buffer. Pressure cook slides for 25 min from the time lock pin slides up, indicating that the unit has pressurized (see Note 8). 2. Cool under running tap water for 30 min (see Note 9). Incubate slides in peroxidase blocking solution for 30 min (see Note 10). 3. Wash with two changes of tap water for 30 s each and transfer to 1 PBS buffer. Use a pap pen to draw a circle around the bladder tissue on slides in order to create a hydrophobic barrier.

3.3.3 Blocking

3.3.4 Primary Antibody Binding

Dry slides carefully around the tissue area with a Kimwipes, adding blocking solution to fully cover the tissue. Incubate in humidity chamber for 30 min at room temperature (see Note 11). 1. Decant off blocking solution by tipping slide along its long edge against a paper towel, and again dry the edges with a Kimwipes. Store in 1 PBS while preparing the dilution of primary antibodies. 2. Make a dilution of primary target antibody (Cleaved Notch1 1:500) (see Note 12) and primary mask antibody (Pan-Cytokeratin 1:500) in 1 PBS-T/BSA antibody buffer. 3. Apply the diluted primary antibodies covering the tissue area. Incubate in humidity chamber overnight at 4  C. Decant off the primary antibodies. Wash slides twice in 1 PBS-T for 10 min each and once in 1 PBS for another 10 min.

3.3.5 Secondary Antibody Binding

1. Dry slides with fresh Kimwipes and apply goat anti-mouse Alexa 546 solution (1:500) in DAKO EnVision™ goat antirabbit (neat) to slides. Incubate in humidity chamber for 1 h at room temperature. 2. Decant off secondary antibodies. Wash slides twice in 1 PBS-T for 10 min each and once in 1 PBS for another 10 min. 3. Dry slides around edge with Kimwipes, and store in 1 PBS while preparing the tyramide working solution.

3.3.6 Tyramide Signal Amplification

1. TSA (Tyramide signal amplification) technology use horseradish peroxidase (HRP) to catalyze covalent deposition of fluorophores directly adjacent to the immobilized enzyme. The labeling reaction is quick (less than 10 min) and deposited labels can then be viewed directly using fluorescence microscopy.

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2. Preparation of tyramide working solution: Cy3 tyramide stock solution is diluted 1/100 in amplification buffer. The tyramide working solution has to be prepared at the last moment to avoid early interaction between tyramides and H2O2 and kept protected from direct light to preserve fluorescent dyes. 3. Apply Cy3 tyramide working solution to each slide (0.25 mL/ tissue area). Incubate sections for 5–10 min at room temperature (see Note 13). 4. Wash out TSA solution from slides twice in 1 PBS-T, first for 5 min, then wash a final time in 1 PBS for 5 min. Cover-Slipping

1. Dry slides with Kimwipes, and apply prolong gold antifade reagent with DAPI (50–75 μL) (see Note 14). Be careful not to create bubbles over the tissue. Gently place coverslip over tissue sections and let dry overnight in the complete absence of light. 2. Seal around edges of the coverslip with clear nail polish to further preserve stain and prevent fading (see Note 15).

Imaging

1. Capture images of the immunolabeled sections using a fluorescent microscope with filter sets for DAPI (excitation filter center wavelength/bandwidth: 378/52 nm, emission filter center wavelength/bandwidth: 447/60 nm) and TRITC/ Cy3 (excitation filter center wavelength/bandwidth: 554/23 nm, emission filter center wavelength/bandwidth: 609/54 nm). 2. Image analysis is performed by using the image processing package ImageJ.

4

Notes 1. All animal care and experimental procedures should comply with the EEC/EU guidelines (Directive 86/609/EEC and Directive 2010/63/EU). 2. This step increases the accessibility of fixative to the inner epithelial surface of bladder tissue. 3. Neutral buffered formalin (NBF) allows thin sectioning, by hardening tissue, preserving the structure and the integrity of bladder epithelium [13]. To achieve efficient penetration of fixative to the epithelial tissue, bladder should be fixed in a sufficient volume of solution; optimally in a ratio of 10:1 fixative to specimen. 4. Long periods of fixation and long periods between the cut section and the staining can lead to high levels of autofluorescence. Immediate fixation is critical.

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For mouse bladder, the optimal fixation time varies between 6 and 24 h(s), depending on the tissue size. A fixation step that lasts longer than 48 h is not recommended, as over fixation may present difficulties in antigen retrieval. 5. Fixing at 4  C, slows down metabolic processes prior to complete fixation. 6. 37% formaldehyde and xylene are carcinogenic. Use always in hood and make sure to use appropriate personal protective equipment when handling the solution or slides that have been treated with these solutions. 7. Drying of sections may result in nonspecific labeling. Antibodies may nonspecifically attach to dried tissue as a result of local ionic charges. Avoid processing too many slides at once in between steps without applying the next appropriate reagent/ solution in order to prevent tissue drying. 8. The antigen retrieval step is critical as it breaks the protein cross-links formed by formalin fixation, and thereby uncovers hidden antigenic sites. Use a sufficient volume of antigen retrieval solution in order to cover the slides by at least a few centimeters and do not allow the slides to dry out from evaporation during the boil. We suggest an antigen retrieval time of 25 min. Less than 25 min may leave the antigens un-retrieved, leading to weak staining. More than 25 min may lead to nonspecific background staining or to sections dissociation from the slides. 9. The gradient cooling allows the antigenic site to re-form after being exposed to high temperature. 10. Some cells or tissues contain endogenous peroxidases. Activation and covalent binding of TSA reagent is catalyzed by peroxidase. Therefore, tyramide reagent can interact with endogenous peroxidases generating nonspecific background signal. Preincubation with saturating amounts of H2O2 irreversibly blocks endogenous peroxidase activity and reduces background. However, certain tissues, cells, or antigens (especially cell surface proteins) can be damaged by high concentrations of H2O2. We therefore recommend 0.3% or 1% for routine applications. 11. Serum blocking is considered more efficient if chosen from the same host species as the secondary antibody. Since our secondary anti target antibody is a goat anti-rabbit antibody, we used normal goat serum in our blocking solution. 12. Using TSA kit, concentration of the primary antibody should be reduced approximately tenfold compared to the concentration used for conventional immunofluorescence

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(IF) detection. We use the cleaved Notch1 (Val1744) (D3B8) antibody diluted between 1:50 and 1:100 in the conventional IF protocol. 13. Prolonged exposure to Cy3 tyramide working solution can increase background and reduce the signal resolution due to diffusion of activated tyramide away from the epitope. 14. DAPI (40 ,6-diamidino-2-phenylindole) binds strongly to AT-rich regions in DNA and is used as a nuclear counterstain. When bound to double stranded DNA, DAPI has a maximum emission wavelength at 461 nm. 15. For long-term storage of immunostained sections, slides can be kept at 20  C. References 1. Kopan R, Ilagan MX (2009) The canonical Notch signaling pathway: unfolding the activation mechanism. Cell 137:216–233 2. Brou C, Logeat F, Gupta N, Bessia C, LeBail O, Doedens JR, Cumano A, Roux P, Black RA, Israel A (2000) A novel proteolytic cleavage involved in Notch signaling: the role of the disintegrin-metalloprotease TACE. Mol Cell 5:207–216 3. Yang G, Zhou R, Zhou Q, Guo X, Yan C, Ke M, Lei J, Shi Y (2019) Structural basis of Notch recognition by human gammasecretase. Nature 565:192–197 4. Petcherski AG, Kimble J (2000) Mastermind is a putative activator for Notch. Curr Biol 10: R471–R473 5. Jarriault S, Brou C, Logeat F, Schroeter EH, Kopan R, Israel A (1995) Signalling downstream of activated mammalian Notch. Nature 377:355–358 6. van Tetering G, Vooijs M (2011) Proteolytic cleavage of Notch: "HIT and RUN". Curr Mol Med 11:255–269 7. Kovall RA, Gebelein B, Sprinzak D, Kopan R (2017) The canonical Notch signaling pathway: structural and biochemical insights into shape, sugar, and force. Dev Cell 41:228–241 8. Rampias T, Vgenopoulou P, Avgeris M, Polyzos A, Stravodimos K, Valavanis C,

Scorilas A, Klinakis A (2014) A new tumor suppressor role for the Notch pathway in bladder cancer. Nat Med 20:1199–1205 9. Faget L, Hnasko TS (2015) Tyramide signal amplification for immunofluorescent enhancement. Methods Mol Biol 1318:161–172 10. Lobry C, Oh P, Aifantis I (2011) Oncogenic and tumor suppressor functions of Notch in cancer: it’s NOTCH what you think. J Exp Med 208:1931–1935 11. Rettig EM, Chung CH, Bishop JA, Howard JD, Sharma R, Li RJ, Douville C, Karchin R, Izumchenko E, Sidransky D, Koch W, Califano J, Agrawal N, Fakhry C (2015) Cleaved NOTCH1 expression pattern in head and neck squamous cell carcinoma is associated with NOTCH1 mutation, HPV status, and high-risk features. Cancer Prev Res (Phila) 8:287–295 12. Stack EC, Wang C, Roman KA, Hoyt CC (2014) Multiplexed immunohistochemistry, imaging, and quantitation: a review, with an assessment of Tyramide signal amplification, multispectral imaging and multiplex analysis. Methods 70:46–58 13. Grizzle WE (2009) Special symposium: fixation and tissue processing models. Biotech Histochem 84:185–193

Methods in Molecular Biology (2021) 2346: 135–149 DOI 10.1007/7651_2020_297 © Springer Science+Business Media New York 2020 Published online: 14 July 2020

Examining Local Cell-to-Cell Signalling in the Kidney Using ATP Biosensing Gareth W. Price, Joe A. Potter, Bethany M. Williams, Chelsy L. Cliff, Mark J. Wall, Claire E. Hills, and Paul E. Squires Abstract Cell-to-cell communication is an essential process for the efficient function of cells and tissues. Central to this is the purinergic transmission of purines, with ligands such as adenosine triphosphate (ATP). Altered cell-to-cell communication, and in particular changes in the paracrine release of extracellular ATP, plays crucial roles in pathophysiological conditions, such as diabetes. ATP biosensing provides a reliable, realtime measurement of local extracellular ATP concentrations. This allows the detection of altered ATP release, which underlies the progression of inflammation and fibrosis and is a potential therapeutic target. Here we describe in a step-by-step basis how to utilize sensitive microelectrode biosensors to detect low, real-time concentrations of ATP, in vitro. Keywords Adenosine triphosphate, Adhesion, Biosensing, Cell-to-cell communication, Connexins, Purines

1

Introduction While purines (adenosine triphosphate, ATP; adenosine diphosphate, ADP; and adenosine) play crucial roles in cellular metabolism and energy storage, they are also important extracellular signalling molecules. Signalling with purines plays pivotal roles in normal physiology, and dysregulation of purinergic signalling can lead to a number of disease states [1, 2]. Thus, real-time measurement of local extracellular purine concentrations provides great value in furthering our understanding of the functional contributions of purines and can also help determine their potential as drug targets when the dysregulation of signalling occurs. Although several techniques have been used to detect ATP and other purines of interest, such as microdialysis, bioluminescent luciferase assays, and “purinoreceptor patch sniffing,” the recent development of microelectrode biosensors represents a step change in technology. Microelectrode biosensors offer a real-time, sensitive method to detect low concentrations of ATP, adenosine (and indeed several other analytes) both in vivo and in vitro [3].

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As summarized by Dale et al. in their review [3], there are several desirable measurement characteristics of any system attempting to detect ATP, all of which biosensors are capable of: l

Temporal resolution: measurements should be obtained within meaningful timescales. ATP release generally occurs on secondminute timescales with the response time of microelectrode biosensors sub-second [4].

l

Analyte resolution: sensitivity within normal physiological concentrations, e.g., micro- to millimolar ranges, and biosensors should show a linear response to increasing analyte concentrations. When optimized, microelectrode biosensors can detect in the nM range [5].

l

Selectivity: for example, ATP biosensors detect ATP but do not detect ADP, AMP, or adenosine. The permselective layer present on the biosensor limits the effects of non-specific electrochemical interferents.

l

Minimally invasive: although more important in an in vivo context, measurements should be taken with minimal disturbance to the cells and tissues.

Here, we describe quantitative characterization of altered cellto-cell communication in the proximal region of the diabetic kidney using ATP biosensing. Diabetic nephropathy represents the leading cause of end-stage renal disease (ESRD) and currently affects one third of people with diabetes (reviewed in [6]). Several structural and functional changes occur of which tubulointerstitial fibrosis is the key underlying pathology [7, 8]. Co-localized at the sites of cell-cell adhesion, transmembrane connexins oligomerize into hexameric assemblies called connexons [9]. When cells are sufficiently coupled, aligned connexons dock to provide a conduit for direct gap junction intercellular communication [10]. We have recently shown in tubular epithelial cells of the proximal nephron that glucose-evoked transforming growth factor-beta1 (TGF-β1) decreases E-cadherin-mediated cell adhesion and consequently gap junction intercellular communication [11, 12]. In the absence of adhesion, connexons form hemichannels through which local signalling mediators such as ATP can be released [10]. In the proximal kidney, ATP has itself been shown to further exacerbate the loss of E-cadherin expression and reduce intercellular ligation forces [13]. The rise in local intercellular ATP is linked to inflammation, fibrosis, and the pathology of a number of diseases [2, 10, 14–16]. Thus, the use of ATP biosensing is an important experimental technique used to resolve the contribution of altered cellto-cell communication in the development and progression of diseases such as diabetic nephropathy.

Biosensing & Hemichannel-Mediated ATP Release

2 2.1

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Materials Tissue Culture

1. Clonal human kidney proximal tubule (HK2) epithelial cell line (ATCC). 2. Tissue culture-grade plastics (T75 flasks, 6-well plates, VWR). 3. Cell culture media: Dulbecco’s modified Eagle medium/ Ham’s F12 nutrient mixture + Glutamine (2 mM) (DMEM/ F-12, Thermo Fisher Scientific), fetal calf serum (FCS, Thermo Fisher Scientific), epidermal growth factor (EGF, SigmaAldrich), penicillin/streptomycin (Sigma-Aldrich). 4. Round 10 mm glass coverslips, thickness #0 (VWR). 5. Glycerol.

2.2

Instrumentation

1. ATP biosensor and null sensors (2 mm length, 50 μm diameter) (Sarissa Biomedical; see Note 1 and Subheading 2.4). 2. Biosensor holders, built in-house following Sarissa Biomedical instructions (see Note 2). 3. Silver/silver chloride reference electrode (Sarissa Biomedical). 4. M-152 Narishige manipulators (Digitimer; see Note 3).

and

magnetic

stands

5. Four-channel desktop 8100-K4 potentiostat and connecting wires (BNC-bulldog clips) (Pinnacle Technology Inc.; see Note 4). 6. Cambridge Electronic Design (CED) Micro1403 AnalogueDigital Board (see Note 5). 7. PC with Spike2 software and Sirenia software. 8. BNC wiring (RS Components). 9. ALA HPT-2A temperature-controlled perfusion tube, TS100 temperature sensor, and TS-10 controller (Scientifica). 10. Custom perfusion chamber (see Note 6). 11. Peristaltic vacuum pumps and tubing (1.6bore, 1.6 mm wall) (Watson-Marlow; see Note 7). 12. KL 1600 LED light source and arm (VWR). 13. Faraday cage (this should be adequately earthed; see Note 8). 14. Metal baseplate (this should be adequately earthed; see Note 9). 15. Articulated stereo microscope (Olympus). 16. Unstirred, heated water bath (Grant Instruments Ltd).

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17. Watchmaker fine tweezers (Amazon). 18. Humidified incubator: 37  C, pH 7.4, 5% CO2. 2.3 Balanced Salt Solution (BSS)

1. Calcium-containing BSS (pH 7.4): Sodium chloride (NaCl, 137 mM), potassium chloride (KCl, 5.36 mM), magnesium sulfate heptahydrate (MgSO47H2O, 0.81 mM), sodium phosphate dibasic dihydrate (Na2HPO42H2O, 0.34 mM), potassium dihydrogen phosphate (KH2PO4, 0.44 mM), sodium bicarbonate (NaHCO3, 4.17 mM), HEPES (4-(2-hydroxyethyl)-1-piperazineethanesulfonic acid, 10 mM), calcium chloride dihydrate (CaCl22H2O, 1.26 mM), glucose (C6H12O6, 2.02 mM). 2. Calcium-free BSS: Same as calcium-containing, with the following changes: Sodium chloride (148 mM), EGTA (ethylene glycol-bis(β-aminoethyl ether)-N,N,N0 ,N0 -tetraacetic acid, 100 μM), zero calcium chloride dihydrate.

2.4

Biosensors

ATP biosensors are platinum-iridium (90%:10%) wires encased in a glass micropipette (Fig. 1). A thin layer of enzymes, namely, glycerol kinase and glycerol-3-phosphate oxidase, are layered on the wire which detect ATP through an enzymatic cascade, ultimately resulting in the production of H2O2 (see Fig. 2 for full cascade). To combat electrochemical interference, the enzyme layer is deposited over a permselective barrier [17–20]. When glycerol is present at concentrations that saturate glycerol kinase (>0.5 mM), the reaction producing H2O2 is directly proportional to the concentration of ATP present. Oxidation of H2O2 occurs at potentials of +500 mV, which can be detected electroanalytically by measuring the current required to maintain the potential within the system (constant potential amperometry). Each experiment requires the use of a null sensor, a microelectrode manufactured in the same way as the ATP-detecting biosensors, but without inclusion of the enzymes. Any change in the null sensor current is a result of non-ATP-derived changes within the system and can be subtracted from the ATP biosensor signal either during or after recording.

Fig. 1 Microelectrode biosensors. A platinum-iridium wire is encased in a glass micropipette. The wire is coated with inner and outer permselective layers, to limit interference from non-selective contaminants, and an enzymatic layer that detects ATP

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A.

C. Current (nA)

B.

2.5 2.0 1.5 1.0 0.5 0.0 0

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Fig. 2 The ATP-detecting glycerol-assisted enzymatic cascade. When glycerol is present in saturating concentrations (>0.5 mM), ATP activates a series of reactions (panel a) which ultimately result in the production of H2O2. This can be detected electroanalytically. Panel (b) shows a representative trace with biosensor responses to different concentrations of ATP. A graph plotting current against ATP concentration is shown in panel (c). ATP biosensors show a linear response to increasing concentrations of ATP. No change in baseline current should be observed with null sensors (data not shown)

3 3.1

Methods Tissue Culture

1. HK2 cells (passages 18–30) were maintained in DMEM/F12 growth media, which has a high basal glucose concentration (17.5 mM) to encourage cell growth. 2. HK2 cells were seeded onto sterile 10 mm #0 glass coverslips in a 6-well plate at a seeding density of 2  104 (see Note 10). 3. Prior to treatment, HK2 cells were cultured in low (5 mM) glucose DMEM/F12 medium for a period of 48 h, followed by a 24 h incubation in serum-free low (5 mM) glucose DMEM/ F12 medium to negate any pre-stimulatory effects from the high (17.5 mM) glucose. 4. After this 24 h period, HK2 cells were treated with a desired stimulus (10 ng/mL TGF-β1) against control HK2 cells for 48 h with a final confluence of 80%.

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Fig. 3 The basic biosensor setup. A perfusion system flows over the cells, which are located in the center of the custom-made chamber. An ATP biosensor and a null biosensor are placed just above the cell monolayer and are connected to an Ag:AgCl reference electrode. A potentiostat and analogue-digital board convert H2O2 into a digital format that can be viewed and analyzed on a computer 3.2 Initial Rig Preparation

To minimize electrical interference, all controllers, pumps, PC, and water bath are located outside of the Faraday cage. Holes are used to allow tubing to enter the center of the cage. The biosensors (and manipulators), perfusing tubes and perfusion chamber, microscope, and lighting tube are the only items inside the cage. A diagram of the below instructions can be found in Fig. 3. 1. Place the Faraday cage and metal baseplate on a secure platform (see Note 11). Set up the microscope inside. The Faraday cage and baseplate should be earthed (using crocodile clips and leads) through the potentiostat (see Note 8). 2. Place the perfusion chamber in the center of the baseplate, and stick down with glue/blue Tac. The chamber can be raised on a metal support if necessary. 3. Place two biosensor holders onto manipulators, and position on either side of the perfusion chamber. Connect the biosensor holders to the potentiostat (channels 1 and 2) via the positive (red) clip from the connecting wires. The potentiostat should be placed outside of the Faraday cage to limit interference. 4. Connect a negative (black) clip from the connection wire to the silver/silver chloride reference, and carefully place inside the perfusion chamber, ensuring that it is submerged (see Note 12). 5. Using BNC wires, connect channels 3 and 4 from the potentiostat to the CED analogue-digital board.

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6. Set up perfusion by using two peristaltic pumps (placed outside of the cage) set to opposing directions. For the input, tubing is threaded through the cage (with the other end placed into the perfusion solution), connected to the temperature-controlled perfusion tube, and then subsequently attached to a needle, which is placed into the right-side compartment of the perfusion chamber. An output tube with another needle is placed in the left compartment, connected to a syringe dropper, and then threaded back out of the cage, through the pump and into a waste beaker. The waste output tube can be placed back into the water beaker to recycle. It is important to keep the level of fluid in the perfusion chamber constant; otherwise, the biosensor signals will oscillate. This can be optimized by changing the flow rate of the two pumps and the positioning of the input and output needles in the recording chamber. 7. Direct the LED light source so that it illuminates the central compartment of the perfusion chamber. Make sure cabling is secure to the baseplate. 3.3 Experiment Preparation

Biosensors detect ATP through an enzymatic cascade that ultimately results in the production of H2O2 and a change in current. Constant potential amperometry allows measurement of ATP concentrations by changing the current required to maintain +500 mV. This analogue data is converted to a digital signal using an analogue to digital (AD) board. The CED Micro1401-3 is a versatile, powerful acquisition unit which offers a 16-bit, 500 kHz maximum aggregate sampling rate and can achieve this with a high degree of precision in real time. The Micro1401-3 is connected to a computer running the acquisition and analytical program, Spike2 (CED). The perfusion solution is pre-warmed (with a water bath) to prevent outgassing and maintained at physiologically relevant temperatures (by the temperature controller). 1. Prepare >1 L balanced salt solution (BSS, calcium-containing and calcium-free), and add 1 mL of glycerol (2 mM) (see Note 13). Take 50–100 mL of the BSS, and make up a calibration solution (10 μM ATP). Place the two solutions and a beaker of dH2O into the pre-warmed water bath (set to 40  C). 2. Before starting experiments perfuse dH2O through the system by turning on both pumps, to clean the tubes and to ensure the bath level is stable. 3. Turn on the temperature controller, initially set the desired temperature to 40  C, and turn the gain and integration to 50% and the limiter to 75–100%. Use the temperature sensor, and alter to the desired temperature, i.e., 37  C media perfusing through the chamber. Once this has been determined, it will remain constant as long as flow rates are unaltered. Make

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Bend

Parallel to coverslip

Fig. 4 A pair of fine forceps bends the biosensor at the white plastic joint so that it lies parallel to the coverslip once inserted into the biosensor holder

sure liquid flows through the temperature-controlled perfusion tube at all times. 4. Turn on the computer and then the potentiostat and analoguedigital board. Open Sirenia and Spike2. 5. Switch the perfusing solution from dH2O to calciumcontaining BSS. After a few minutes, place the waste tube into the BSS container to recycle the BSS. 6. To maximize the detection of any changes in extracellular ATP concentration, the sensing area of the ATP biosensor has to be positioned as close to the cell monolayer as possible. This is achieved by carefully bending the biosensor so the sensing area is parallel to the cell monolayer (which will be placed in the bottom of the perfusion chamber). The best way to do this is to place the biosensor in the holder above the perfusion chamber and focus on it using the microscope. Using fine forceps, carefully bend the biosensor (at the white plastic joint, without touching the sensing area) until it is at the correct angle (see Fig. 4). Before commencing an experiment, check that the sensor can be placed close and parallel to the bottom of the bath. The same process should then be repeated for the null sensor. Again, check that both sensors can be placed close to the bottom of the bath (the manipulators may need to be moved to achieve this). The biosensor and the null sensor should be positioned close together, but not touching, as this would cause interference. 7. If not done already, insert the biosensor and null sensor into their appropriate holders. Carefully lower them into the

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perfusing liquid until they are submerged. Once wet, it is important to keep the sensors in solution most of the time; if they become dry, they will lose sensitivity (see Note 17). 3.4 Biosensor Polarization

Before measuring extracellular ATP release from cell monolayers, the biosensor and null sensor have to be polarized. The biosensor and null sensor act like capacitators; thus, when they are initially held at +500 mV, a current flows to charge them, and this baseline current will exponentially decay over time. This baseline current will never reach zero, but the gradient of the decay in baseline current will get less over time. After 30–60 min, the gradient of the decay in baseline current will be very low, and accurate measurements can now be made. This procedure may need to be repeated multiples times throughout the day when changing coverslips, or if biosensor sensitivity is low and new biosensors have to be used (see Note 14). Sensitivity can be assessed by brief perfusion with 10 μM ATP. Each time the biosensors are removed from solution, they will need time to repolarize. Therefore, if possible, keep the biosensors in the solution at the side of the chamber when you need to change coverslips (in some cases this will not be possible as the coverslip fills the perfusion chamber). 1. In Sirenia, click New Experiment, name the file appropriately, and then click Add Device. Click on the potentiostat from the list, choose the option to use channels 3 and 4 for outputs, and then press ok (see Note 15). 2. Open the Configuration tab and set the potentials to 0.5 V and a max current of 100 μA. Set all gains on all channels. 3. Make sure channels 3 and 4 are set to output. There should not be ticks by their names. 4. Select four samples a second (4 Hz). Click Accept and then the red record button. 5. In Spike 2, start a new data document. Configure the program to receive two inputs, name them appropriately (NULL and ATP), and set their units to nA by clicking Configuration in the menu bar. A virtual channel (Channel->New Virtual Channel) can be used with the expression Ch(2)-Ch(1) to subtract the null from the ATP readings online. It is recommended to smooth the traces over a 1–2 s duration by right clicking each trace and clicking Channel Process. Press Start in the toolbar to start recording (see Note 16). 6. The current in the ATP and NULL channels should resemble a negative exponential, asymptotic to the horizontal axis. Once a stable baseline occurs, press the Stop button in Spike 2 and Sirenia, save the file in Spike, and leave the biosensors in the perfusion chamber until ready to proceed with the next section, ensuring that perfusing media continues to recycle properly (see Note 17).

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Experiment

1. Carefully raise the biosensors out of the perfusing media, and place the coverslip into the central compartment of the perfusion chamber. Lower the biosensors until they lie immediately above the cell monolayer (see Note 18). 2. Start new files in Sirenia and Spike2, with the same configurations used to polarize the biosensors; however, use a max current value of 25 nA to increase resolution (see Note 19). 3. Once the new files are created, allow calcium-containing BSS to perfuse over the cells until a stable baseline occurs (normally 5–10 min) which allows cellular climatization. 4. Switch the perfusing media to calcium-free BSS (see Note 20), and allow sufficient time for a response to occur (see Note 21). 5. Switch back to calcium-containing BSS. 6. After approximately 5 min, lift the biosensors so they are above the cell layer but still in the perfusing medium. Then perfuse with the 10 μM ATP solution for a calibration trace, ensuring that the waste output is not recycled. Make sure the calibration trace has reached steady state before washing with normal calcium-containing BSS. If not using a new biosensor or if you are unsure about the sensitivity of the biosensor, the calibration step can be carried out before the experiment. This way you will prevent the waste of time and reagents if the biosensor has such low sensitivity that no ATP is detected.

3.6

Analysis

Spike 2 is a powerful analysis tool that can be used extensively to analyze traces and can help to remove undesirable noise and artefacts. Simple analysis of ATP biosensor traces to achieve approximate quantification of changes in ATP concentrations is outlined below. 1. Bring up the cursors (ctrl-1 and ctrl-2), and align them to the start and end of the major ATP peak that is selected for analysis (Fig. 5). 2. Display Y values corresponding to each cursor point by clicking Cursor ! Display Y values (Fig. 6). A panel will appear with the Time (x-axis) and Current (y-axis, nA), note the Y values, and calculate their difference (see Note 22). 3. Calculate the amplitude of the 10 μM ATP calibration peak by repeating step 2. 4. Absolute concentrations can be calculated from comparing the amplitude of the selected peaks to that of the ATP calibration peak. For example, a 1 nA peak corresponds to 5 μM if the ATP calibration peak is 2 nA. 5. Can repeat this but after subtracting the null channel. 6. After the desired number of replicates, results can be statistically tested using an ANOVA.

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Fig. 5 An example of how to place cursors around an ATP peak

Fig. 6 An example of the dialogue box which contains Y values for cursor thresholds

4

Notes 1. Although shorter and thinner biosensors are available, the 2 mm/50 μM biosensor provides maximum detection area, increasing sensitivity and reliability. 2. Sensor holders were made following Sarissa Biomedical instructions, found at http://www.sarissa-biomedical.com/ how-to-use/technical-tips/sensor-holder.aspx. To make your own sensor holders, you will need a cable (thinner than a 1 mL syringe), a 1 mL syringe, a gold-plated socket for a 1 mm diameter pin (buy from Sarissa), and some superglue and soldering kit. 3. While more sensitive or motorized micromanipulators are available from Narishige or other suppliers, these course, manual manipulators are sufficient to position the biosensors correctly and are reasonably priced. 4. This protocol uses four channels: two inputs for ATP biosensor and null sensors and the other two as outputs to transfer

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analogue data from the potentiostat into the analogue-digital board. An analogue potentiostat such as that purchasable on Sarissa Biomedical/Whistonbrook would only need two channels. 5. Even though the Pinnacle potentiostat possesses an onboard analogue-digital board, the external Cambridge Electronic Design system (analogue-digital board and Spike2 software) allows greater experimental control, including online analysis and rigorous data processing. 6. Although here we use an in-house custom chamber consisting of pipette sections glued onto a large Petri dish, commercial chambers are available, such as those from Warner Instruments. A small metal (or plastic) mesh can often be useful to raise coverslips, ease recovery of coverslips post-experiment, and allow perfusion underneath the coverslips. 7. The internal diameter will affect speed of perfusion over cells. Faster perfusion speeds will decrease the lag period and increase the rate of onset for stimuli. However, this will also increase the likelihood of frequent volume changes, which will result in noise in recording traces, so a balance needs to be achieved. If required, coverslips can be carefully broken so they fit in the bath. 8. The use of an earthed, metal Faraday cage and baseplate ensures that external electrical interference is kept to a minimum. If artefacts or noise are common during recording, then earthing is an important factor to check. If cost is a consideration, then a simple Faraday cage can be constructed using a large cardboard box covered in foil or using wire mesh attached to plastic greenhouse supports. 9. The metal baseplate provides an area where the manipulators can be secured but also contributes to proper earthing. Any thickness can be used, but it is best if it is magnetic; however, ensure the size covers the inside of Faraday cage. 10. Seeding density is an important variable that requires initial optimization by the user. Cells seeded too sparsely release extracellular ATP at a concentration below biosensor detection levels. Too many cells could alter cell viability and artificially increase release of ATP through cell rupture obscuring physiological responses. 11. If unsecure, vibrations will cause noise during readings. If available, an antivibration table could be used to dampen vibrations and improve reliability. Alternatively, the baseplate can be placed on partially inflated inter-tubes or tyres. 12. Make sure that the negative connection does not contact a positive. Placing the black clip into a separate Petri dish (stuck down using glue/blue Tac) allows proper isolation.

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13. Although enough glycerol must be added in saturating concentrations, ensure the final concentration is kept constant between experiments. Adding excessive glycerol may cause unstable and inconsistent readings. Failing to add glycerol prevents the biosensor from detecting ATP (as it is an enzyme co-factor), which will be visualized as a lack of response to added ATP during calibration. This can be used as a control, showing that any currents measured are absent without glycerol confirming that they represent changes in ATP. 14. If polarization fails to regain sensitivity, cycling the biosensors from +500 mV to 500 mV at 100 mV/s for 10 cycles may help. This can be achieved through the “Conditioning” button in the configuration panel of the Sirenia interface. 15. This initial screen often does not save the configuration to the device; however, it is useful to set the outputs so the software displays information correctly. If the device fails to appear in the list, check all connections, and make sure all hubs and equipment are powered on. 16. If a static signal is observed in Spike 2, this could be a failure of the potentiostat to output the analogue data correctly. Restarting the experiment in Sirenia is usually a quick fix. 17. Once wet, ensure biosensors are always kept in liquid, either in the perfusion chamber or a separate pot (also available from Sarissa Biomedical) filled with balanced salt solution. Allowing biosensors to dry may irreversibly damage the biosensors. Multiple successive removals of biosensors from the perfusion chamber (to allow placing of new coverslips) may result in the loss of sensitivity, after which polarization is required. 18. The distance between the biosensor and the cell layer is an important variable. Reduction in biosensor current amplitude will occur if the sensing area of the biosensor is at too great a distance from the cell layer (as the released ATP will dissipate); conversely, if the biosensor is positioned too close to the cell monolayer, it may damage cells (releasing ATP) or cause damage to the biosensor. It is helpful to establish the correct biosensor position required without cells being present, noting the manipulator measurements and then returning to this state once the coverslip is placed underneath. 19. Normally 25 nA is more than sufficient for the majority of physiological responses and maximizes sensitivity and resolution. However, this value may need to be changed for other experiments where ATP concentrations are higher or sensor sensitivity is especially high. 20. To ease analysis, use text comments (ctrl-T) to mark key events in the experiment.

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21. Depending on experimental conditions (such as cell type, treatment, or hemichannel opening stimuli), the cells may require a preload with ATP to ensure that concentrations are sufficiently detectable by the biosensor [11, 21]. This can be achieved by placing the coverslip into calcium-free BSS for 5 min before adding ATP (10 mM) in calcium-free BSS for 10 min and finally 10 min washing in calcium-containing BSS. 22. Alternatively, you can hover the mouse cursor over the start and end of each peak, and their values should appear in the tooltip.

Acknowledgments The authors acknowledge the generous support of Diabetes UK (PES & CEH: 16/0005427, 16/0005544, and 18/0005919), the Royal Society (CEH), and an EFSD/Boehringer Kidney Award (CEH & PES). References 1. Burnstock G (2018) The therapeutic potential of purinergic signalling. Biochem Pharmacol 151:157–165. https://doi.org/10.1016/j. bcp.2017.07.016 2. Prakoura N, Kavvadas P, Chadjichristos CE (2018) Connexin 43: a new therapeutic target against chronic kidney disease. Cell Physiol Biochem 49:998–1009. https://doi.org/10. 1159/000493230 3. Dale N, Frenguelli BG (2012) Measurement of purine release with microelectrode biosensors. Purinergic Signal 8(27):40. https://doi.org/ 10.1007/s11302-011-9273-4 4. Wall MJ, Richardson MJE (2015) Localized adenosine signaling provides fine-tuned negative feedback over a wide dynamic range of neocortical network activities. J Neurophysiol 113:871–882. https://doi.org/10.1152/jn. 00620.2014 5. Wall MJ, Dale N (2013) Neuronal transporter and astrocytic ATP exocytosis underlie activitydependent adenosine release in the hippocampus. J Physiol 591:3853–3871. https://doi. org/10.1113/jphysiol.2013.253450 6. Price GW, Potter PA, Williams BM et al (2020) Connexin-mediated cell communication in the kidney: a potential therapeutic target for future intervention of diabetic kidney disease? Joan Mott Prize Lecture. Exp Physiol 105:219–229. https://doi.org/10.1113/ EP087770

7. Humphreys BD (2018) Mechanisms of renal fibrosis. Annu Rev Physiol 80:309–326. https://doi.org/10.1146/annurev-physiol022516-034227 8. Liu B-C, Tang T-T, Lv L-L, Lan H-Y (2018) Renal tubule injury: a driving force toward chronic kidney disease. Kidney Int 93:568–579. https://doi.org/10.1016/j. kint.2017.09.033 9. Bosco D, Haefliger J-A, Meda P (2011) Connexins: key mediators of endocrine function. Physiol Rev 91(1393):1445. https://doi.org/ 10.1152/physrev.00027.2010 10. Hills CE, Price GW, Squires PE (2015) Mind the gap: connexins and cell–cell communication in the diabetic kidney. Diabetologia 58:233–241. https://doi.org/10.1007/ s00125-014-3427-1 11. Hills C, Price GW, Wall MJ et al (2018) Transforming growth factor Beta 1 drives a switch in Connexin mediated cell-to-cell communication in tubular cells of the diabetic kidney. Cell Physiol Biochem 45:2369–2388. https://doi.org/10.1159/000488185 12. Siamantouras E, Hills CE, Squires PE, Liu K-K (2016) Quantifying cellular mechanics and adhesion in renal tubular injury using single cell force spectroscopy. Nanomedicine 12:1013–1021. https://doi.org/10.1016/j. nano.2015.12.362

Biosensing & Hemichannel-Mediated ATP Release 13. Siamantouras E, Price GW, Potter JA et al (2019) Purinergic receptor (P2X7) activation reduces cell–cell adhesion between tubular epithelial cells of the proximal kidney. Nanomedicine 22:102108. https://doi.org/10.1016/j. nano.2019.102108 14. Mugisho OO, Green CR, Zhang J et al (2019) Connexin43 hemichannels: a potential drug target for the treatment of diabetic retinopathy. Drug Discov Today 24:1627–1636. https:// doi.org/10.1016/j.drudis.2019.01.011 15. Burnstock G, Knight GE (2018) The potential of P2X7 receptors as a therapeutic target, including inflammation and tumour progression. Purinergic Signal 14:1–18. https://doi. org/10.1007/s11302-017-9593-0 16. Price GW Chadjichristos CE, Kavvadas P et al (2020) Blocking connexin-43 mediated hemichannel activity protects against early tubular injury in experimental chronic kidney disease. Cell Commun Signal 18:79. https://doi.org/ 10.1186/s12964-020-00558-1 17. Calia G, Rocchitta G, Migheli R et al (2009) Biotelemetric monitoring of brain neurochemistry in conscious rats using microsensors and

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biosensors. Sensors 9:2511–2523. https://doi. org/10.3390/s90402511 18. Chirizzi D, Malitesta C (2011) Potentiometric urea biosensor based on urease immobilized by an electrosynthesized poly (o-phenylenediamine) film with buffering capability. Sens Actuators B 157:211–215. https://doi.org/10.1016/j.snb.2011.03.051 19. Dash MB, Douglas CL, Vyazovskiy VV et al (2009) Long-term homeostasis of extracellular glutamate in the rat cerebral cortex across sleep and waking states. J Neurosci 29:620–629. https://doi.org/10.1523/jneurosci.5486-08. 2009 20. Tian F, Gourine AV, Huckstepp RTR, Dale N (2009) A microelectrode biosensor for real time monitoring of L-glutamate release. Anal Chim Acta 645:86–91. https://doi.org/10. 1016/j.aca.2009.04.048 21. Huckstepp RTR, Eason R, Sachdev A (2010) Dale N CO2-dependent opening of connexin 26 and related β connexins. J Physiol 588:3921–3931. https://doi.org/10.1113/ jphysiol.2010.192096

Methods in Molecular Biology (2021) 2346: 151–164 DOI 10.1007/7651_2020_338 © Springer Science+Business Media New York 2020 Published online: 15 December 2020

Isolation and Assessment of Pancreatic Islets Versus Dispersed Beta Cells: A Straightforward Approach to Examine Cell–Cell Communication Rachel T. Scarl, William J. Koch, Kathryn L. Corbin, and Craig S. Nunemaker Abstract Islets of Langerhans, found in the pancreas, are microorgans essential for glucose homeostasis within the body. Many cells are found with an islet, such as beta cells (~70%), alpha cells (~20%), delta cells (~5%), F cells (~4%), and epsilon cells (1%), each with its own unique function. To better understand the roles of these cells and how cell communication alters their function, several techniques have been established such as islet isolation and beta cell dispersion. Here we describe how to isolate primary rodent islets, disperse pancreatic islets, measure intracellular calcium, and use immunofluorescent staining to distinguish beta cells and alpha cells. Key words Beta cells, Dispersion, Immunofluorescence, Intact, Intracellular calcium, Islets of Langerhans, Pancreas

1

Introduction Islets of Langerhans within the pancreas are essential microorgans involved in regulating glucose homeostasis throughout the body. As is true for many organs, communication among a variety of cells with unique functions must occur in order to maintain optimal physiological homeostasis. Insulin, for example, is tightly regulated and is only released by a coordinated response of the population of beta cells within an islet [1, 2]. This cell-to-cell communication is essential to a full understanding of normal islet physiology and pathophysiology. Over the years, several methods have been established to aid in studying communication among cells within an islet. Connexins, proteins involved in connecting one cell to another while allowing the transfer of signaling molecules between cells, have been knocked out in Cx36 / mice, which has resulted in disruptions

Rachel T. Scarl and William J. Koch contributed equally to this work.

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in pulsatility, insulin release, and glucose homeostasis [3]. Paracrine and autocrine signaling though hormones and other secreted factors is another important method of cell–cell communication within the islet [4]. Another interesting theory is that of “hub cells” or cells thought to dictate and control the actions of the surrounding cells. Hub cell ablation has shown a loss of calcium oscillation and loss of regulated insulin release within an islet [5, 6]. Organotypic culture has become a useful technique to not only study how islet cells function almost in vivo but also how other surrounding cells are essential to produce a synchronized response [7, 8]. Another method to allow the researcher to isolate islet cells from any form of cell-cell communication is by dispersing the islets into single cell culture preparations. This prevents gap junction coupling and electrical depolarization or calcium waves to travel from cell to cell and dilutes paracrine chemical signals by increasing the distance between cells. Using live-cell imaging, we have made direct comparisons between intact islets and dispersed islet cells to identify differences in metabolic oscillations [9] and to show that dispersed cells lose the imprinted oscillatory signature of their in vivo environment [10]. We have recently shown that it is possible to reconstruct the physiological responses of dissociated cells from the intact islet, thus deepening our understanding of the role of individual islet cells [11]. In this chapter, we describe the steps to study both intact islets and individual islet cells to better understand each cell’s unique role in a largely coordinated response.

2 2.1

Materials Islet Isolation

1. Thumb dressing and London college forceps. 2. Mayo and sharp-pointed dissecting scissors. 3. Johns Hopkins Bulldog clamp. 4. 5 mL Luer-lock syringe with 30-gauge 0.5-in. needle. 5. 1 mL Luer-lock syringe with 25-gauge needle. 6. 2-in. by 2-in. gauze. 7. 40 (420 μm) sieve (Bellco Glass Inc., NJ). 8. 10 mL pipettes. 9. 15 mL and 50 mL conical tubes. 10. 10 mm petri dishes. 11. 37  C water bath. 12. Weighing scale. 13. Centrifuge (290–453  g). 14. CO2 gas for animal euthanasia (see Note 1).

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15. 70% isopropyl alcohol. 16. G-solution (see Note 2). 17. Collagenase Solution (see Note 3). 18. Histopaque 1100 Solution (see Note 4). 19. RPMI 1640 + L-Glutamine (Invitrogen, CA, Cat: 11875-093) (see Note 5). 20. 10% fetal bovine serum (R&D Systems Inc., MN). 21. 1% penicillin (100 U/L)/streptomycin (100 μL/mL) (Invitrogen, CA, Cat: 15140-122). 22. 37  C humidified atmosphere containing 5% CO2. 2.2

Islet Dispersion

1. Poly-D-lysine (Sigma-Aldrich, MO)–coated glass coverslips (see Note 6). 2. London college forceps. 3. Glass specimen tube and glass pasture pipets coated with Sigmacote (Sigma-Aldrich, MO). 4. Dissociation buffer (see Note 7). 5. RPMI 1640 + L-Glutamine (Invitrogen, CA, Cat: 11875-093) (see Note 5). 6. 10% fetal bovine serum (R&D Systems Inc., MN). 7. 1% penicillin (100 U/L)–streptomycin (100 μL/mL) (Invitrogen, CA, Cat: 15140-122). 8. 37  C humidified atmosphere containing 5% CO2. 9. 6-well plate. 10. 50 mL conical tube. 11. 1000 μL micropipette with 1 mL micropipette tips. 12. Parafilm. 13. 37  C water bath. 14. Trypsin (Invitrogen, CA, Cat: 25300-054). 15. Centrifuge (800 rpm). 16. Aluminum foil. 17. Fume hood.

2.3

Calcium Imaging

1. cellSens Dimension 1.13 imaging software (Olympus, Tokyo, Japan). 2. Fura-2 AM fluorescent dye solution (Invitrogen, CA) (see Note 8). 3. In-line heater at 37  C. 4. Peristaltic pump. 5. Open diamond bath imaging chamber (Warner Instruments, Cat: 64-0288).

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6. Stage adapter (Warner Instruments, Cat: 64-0298). 7. Hamamatsu ORCA-Flash4.0 digital camera (Hamamatsu Photonics K.K., Japan, Model C11440-22CU). 8. BX51WIF fluorescence microscope with 10 objective (Olympus, Tokyo, Japan) (see Note 9). 9. Excitation light via xenon burner supply and filter wheel (Sutter Instrument Co., Novato, CA, Model LB-LS/30). 10. Lambda 10-3 Optical Controller (Sutter Instrument Co., Novato, CA, Model LB10-3-1572). 11. 37  C humidified atmosphere containing 5% CO2. 12. 12-well plate. 13. Desired glucose solution (see Note 10). 14. Aluminum foil. 2.4

Islet Cell Staining

1. 1 PBS (Sigma-Aldrich, MO). 2. 0.5% Triton X100 in 1 PBS. 3. 70% glycerol in 1 PBS. 4. Paraformaldehyde solution, 4% in 1 PBS (Thermo Fisher Scientific, CA). 5. Glass coverslips. 6. Blocking Solution (see Note 11). 7. Anti-insulin antibody guinea pig (Abcam, Cambridge, MA, Cat: ab7842). 8. Goat anti-guinea pig IgG H&L Alexa Fluor 488 antibody (Abcam, Cambridge, MA, Cat: ab150185). 9. Anti-glucagon antibody mouse (Abcam, Cambridge, Cat: ab10988). 10. Goat anti-mouse IgG H&L Alexa Fluor 647 antibody (Abcam, Cambridge, MA, Cat: ab150115). 11. Clear nail polish. 12. 6-well plate. 13. Parafilm. 14. dH2O. 15. Hamamatsu ORCA-Flash4.0 digital camera (Hamamatsu Photonics K.K., Japan, Model C11440-22CU). 16. BX51WIF fluorescence microscope with 10 objective (Olympus, Tokyo, Japan) (see Note 9). 17. Excitation light via xenon burner supply and filter wheel (Sutter Instrument Co., Novato, CA, Model LB-LS/30). 18. Lambda 10-3 Optical Controller (Sutter Instrument Co., Novato, CA, Model LB10-3-1572).

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Methods Islet Isolation

1. Mouse pancreatic islets were isolated using collagenase digestion and gradient centrifugation. For a more in-depth overview and protocol, refer to [12]. 2. Mice were euthanized with CO2 and exsanguinated by heart perfusion using a 1 mL syringe with 25-gauge needle. 3. Mouse abdomen was sterilized with 70% alcohol before cutting open to expose the peritoneal cavity (see Note 12). 4. Finding the common bile duct, a Johns Hopkins Bulldog clamp was used to obstruct the junction between the common bile duct and small intestine. 5. Using a 5 mL syringe with 30-gauge 0.5-in. needle, the pancreas was inflated via the common bile duct with chilled Collagenase Solution. 6. The pancreas was then removed and placed into a 15 mL conical tube containing 1 mL G-Solution for ~8 min at 37  C followed by quick shake for 5 s (see Note 13). 7. Next, the digested pancreas underwent a series of centrifugation and separation steps. The 15 mL conical tube was centrifuged for 2 min at 290  g and the supernatant was discarded. 8. The pellet was then resuspended with 10 mL of G-Solution and centrifuged for 2 min at 290  g. The supernatant was discarded. 9. Again, the pellet was resuspended with 10 mL of G-Solution, and this time filtered using a 40 (420 μm) sieve into a 50 mL conical tube. 10. To resuspend the pellet, 20 mL of G-Solution was added to the 50 mL conical tube and then centrifuged for 2 min at 290  g. The supernatant was discarded. 11. Using 10 mL of Histopaque 1100 Solution, the pellet was resuspended and centrifuged for 20 min at 290  g. This supernatant was transferred to a new 50 mL conical tube with 25 mL of G-Solution and centrifuged for 4 min at 453  g. Then the supernatant was discarded. 12. Using 10 mL of G-Solution, the pellet was resuspended and centrifuged for 3 min at 290  g. The supernatant was discarded, and the pellet was resuspended with culture medium. 13. Islets were incubated overnight (~24 h) in RPMI 1640 + LGlutamine supplemented with 10% fetal bovine serum and 1% penicillin (100 U/L)/streptomycin (100 μL/mL) to allow recovery from collagenase digestion before further treatment (see Note 14).

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Islet Dispersion

1. Autoclave microscope coverslips, London college forceps and glassware before dissociation. Allow sufficient time for materials to dry completely (see Note 15). 2. The day before dispersion, isolate pancreatic islets as described above. It is important to allow an overnight (~24 h) recovery period postisolation. Islets are to be kept at 37  C in a humidified atmosphere containing 5% CO2. 3. Prepare coverslips by placing one slip per well into a 6-well plate. Coat the coverslip with poly-D-lysine solution and place into a fume hood for 15 min. Aspirate the excess liquid off of the coverslips (see Note 16). Keep newly coated coverslips at 37  C in a humidified atmosphere overnight (~24 h) to allow complete drying. 4. In a 37  C water bath, aliquot and warm RPMI 1640 + LGlutamine supplemented with 10% fetal bovine serum and 1% penicillin (100 U/L)/streptomycin (100 μL/mL) in a 15 mL conical tube. Use a minimum of 4 mL per 100 islets dissociated. 5. Also, in a 37  C water bath, aliquot and warm Trypsin in a 1.5 mL Eppendorf tube. Use a minimum of 500 μL per 100 islets dissociated. 6. Pipette Trypsin into a glass specimen tube coated with Sigmacote. Next, pipette isolated islets into the glass specimen tube. 7. Using the pasture pipette coated with Sigmacote, rapidly pipette the solution against the side of the specimen tube while trying to avoid excessive bubbling or suctioning into the pipette (see Note 17). 8. Add 1.5 mL prewarmed RPMI 1640 media to the specimen tube containing dissociated islets. This will neutralize the action of the trypsin. 9. Cover the specimen tube with parafilm and centrifuge the cell suspension at 800 rpm for 3 min. 10. Carefully remove the supernatant without disrupting the cell pellet (see Note 18). Resuspend the pellet in warmed RPMI media (500 μL per 100 islets dissociated). 11. Using the pasture pipet coated with Sigmacote, pasture pipet the cell suspension onto the poly-D-lysine–coated coverslips previously prepared in the 6-well plate (see Note 19). 12. Place the 6-well plate containing the poly-D-lysine coverslips with newly added cell suspension into a 37  C humidified atmosphere containing 5% CO2 for 45 min to 1 h. This allows for cells to settle and attach.

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Fig. 1 Bright field images from dispersed beta cells vs. intact islets. Representative images of individual islet cells (a) and intact islets (b) from mice. Islets typically range from 100 to 200 μm in diameter and contain ~500–2000 cells. Inset in (a) provides higher magnification of three islet cells within the box

13. Following incubation, carefully add 2 mL of fresh prewarmed RPMI 1640 medium to each well while minimizing the disturbance of the cells. 14. Keep dispersed cells in a 37  C humidified atmosphere containing 5% CO2 overnight (~24 h) to allow for recovery before further treatment. 15. See Fig. 1 for brightfield image comparison of intact islets versus dispersed islet cells. 3.3

Calcium Imaging

1. Much equipment is required in order to perform calcium imaging as stated in the materials. 2. Using a peristaltic pump, pass dH2O through an inline heater set at 37  C into a diamond bath imaging chamber mounted to a stage adaptor on a BX51WIF fluorescence microscope with a 10 objective (see Note 20). 3. Next, in a 12-well plate, add 1 mL desired glucose solution and 1 μL Fura-2 AM fluorescent dye solution per well. Once prepared, warm the plate in a 37  C humidified atmosphere containing 5% CO2. 4. In the warmed 12-well plate, add 20–25 intact islets to the Fura-2 AM prepared well. Cover with foil to block out light and place into a 37  C humidified atmosphere containing 5% CO2 for 30 min.

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5. For dispersed islets, in a similar way, add the coverslip with dispersed cells into a prewarmed Fura-2 AM prepared 12-well plate and place into a 37  C humidified atmosphere containing 5% CO2 for 30 min. 6. While cells are being incubated, change the peristaltic pump so that it is passing the desired glucose solution through the inline heater into the diamond bath imaging chamber rather than the dH2O. 7. After incubation, place islets or coverslip with dispersed cells into the diamond bath imaging chamber mounted on the microscope. 8. Observation of the islets or dispersed cells was performed using a Hamamatsu ORCA-Flash4.0 digital camera mounted to a BX51WIF fluorescence microscope. 9. Excitation light was provided using a xenon burner supplied to the image field through a light pipe and filter wheel with a Lambda 10-3 Optical Controller. 10. Regions of interest were drawn around each islet or dispersed cell in order to measure fluorescence intensity per islet or per dispersed cell (see Note 21). 11. Sequential images are taken with 340 nm and 380 nm excitation to produce an intracellular calcium ratio from emitted light at 510 nm. 12. Using cellSens Dimension 1.13 imaging software, the intracellular calcium changes of intact islets and dispersed cells were measured and analyzed (Fig. 2). 3.4

Islet Cell Staining

1. For dispersed islets, follow dispersion protocol in Methods above including fixing dispersed cells to poly-D-lysine–coated coverslip. Once fixed and incubated overnight (~24 h) in RPMI 1640 media, coverslips were washed with 1 PBS and fixed for staining using paraformaldehyde solution. Paraformaldehyde fixed coverslips were then kept at room temperature for 20 min. 2. Following fixation, wash coverslips four times with 1 mL 1 PBS (see Note 22). 3. Next, permeabilize the cells by placing the coverslips in a 6-well plate containing 0.5% Triton X100 in 1 PBS for 10 min at room temperature. Wrap samples with parafilm in order to prevent evaporation. 4. Transfer the permeabilized cells into blocking solution for 1 h. Following incubation, add appropriate primary and secondary antibody.

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Fig. 2 Calcium recordings from dispersed beta cells vs. intact islets. (a, b) Representative examples of intracellular calcium traces using fura-2 AM from individual beta cells (a) or intact islets (b) during sequential stimulation with 4, 10, and 16 mM glucose. Dotted vertical lines indicate changes in glucose. Scale bars indicate the fura-2 AM ratio (340/380 nm) and time. Note the larger changes in calcium and variety of patterns from individual beta cells compared to intact islets. (c, d) Mean calcium from the three representative traces for beta cells (c) and islets (d). Variability is much greater for beta cells as indicated by error bars, but the combined pattern from three different beta-cell traces is closer to resembling the calcium response of an intact islet. The combined patterns of many additional beta cells more closely resemble the response of an intact islet. (For additional details, refer to Scarl et al., Cell Calcium, 2019)

5. For insulin staining, add guinea pig anti-insulin antibody (1:200 dilution) for 2 h at room temperature or keep overnight at 4  C. Then, wash coverslips four times with 0.5% Triton X100 in 1 PBS at room temperature. Next, add goat antiguinea pig antibody for 1 h at room temperature. Wash coverslips four times with 0.5% Triton X100 in 1 PBS.

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6. For glucagon staining, add mouse anti-glucagon antibody (1:200 dilution) for 2 h at room temperature or keep overnight at 4  C. Then, wash coverslips four times with 0.5% Triton X100 in 1 PBS at room temperature. Next, add goat antimouse antibody for 1 h at room temperature. Wash coverslips four times with 0.5% Triton X100 in 1 PBS. 7. Mount stained coverslips to using 70% glycerol in 1 PBS by adding 5–10 μL per coverslip. Then seal coverslips using clear nail polish. Once dry, rinse with dH2O. 8. Observation of stained islets or dispersed cells was performed using a Hamamatsu ORCA-Flash4.0 digital camera mounted to a BX51WIF fluorescence microscope. 9. Excitation light was provided using a xenon burner supplied to the image field through a light pipe and filter wheel with a Lambda 10-3 Optical Controller. 10. Using cellSens Dimension 1.13 imaging software, fluorescent images were taken with 488 nm excitation/525 nm emission for dispersed beta cells (insulin, green fluorescence) and 535 nm excitation/620 nm emission for dispersed alpha cells (glucagon, red fluorescence) (Fig. 3).

Fig. 3 Fluorescent images of dispersed beta cells vs. dispersed alpha cells. Representative fluorescent images of individual beta cells (a) and individual alpha cells (b) found within a single set of dispersed islets. Green fluorescence is indicative of insulin found in beta cells (a). Red fluorescence is indicative of glucagon found in alpha-cells (b). The proportion of beta-cells and alpha-cells in this dispersion accurately represent the distribution of islet cells within an intact islet. The majority of cells found within an intact rodent islet are beta cells (~70%) whereas alpha cells are the second most populous cell type (~20%). Images above depict same field of view

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Notes 1. When conducting animal research, investigators require proper training and approval from the institutional Animal Care and Use Committees in accordance with accepted standards of humane animal care. 2. G-solution is prepared with 1 HBSS (Invitrogen, CA) + 0.35 g NaHCO3/L + 1% bovine serum albumin (BSA). 3. Collagenase solution is prepared with 1.4 mg/mL Collagenase-P (Roche Diagnostics, IN) in G-Solution. 4. Histopaque 1100 Solution is prepared with 100 mL Histopaque 1077 (Sigma-Aldrich, MO) + 120 mL Histopaque 1119 (Sigma-Aldrich, MO). After preparing the Histopaque solution, it is important to keep solution in the dark for optimal use. 5. 500 mL RPMI 1640 + L-glutamine (Invitrogen, CA) is prepared with 50 mL FBS (R&D Systems Inc., MN) + 5 mL penicillin (100 U/mL)–streptomycin (100 μg/mL) (Invitrogen, CA). Media should be stored at 4  C. 6. Poly-D-lysine (Sigma-Aldrich, MO, Cat: P7280-5MG) is prepared by adding 50 mL dH2O to the 5 mg poly-D-lysine bottle using sterile technique. 7. Dissociation buffer is prepared with 0.5 g/L of trypsin (Invitrogen, CA) and 0.2 g/L of EDTA·4Na in Hanks’ Balanced Salt Solution without CaCl2, MgCl2 and MgSO4. 8. Stock solution of fura-2 AM fluorescent dye is prepared with 45 μL DMSO and 5 μL pluronic acid per 1 50 μg vial. For use, add 1 μL Fura-2 AM stock solution to 1 mL glucose solution (1:1000 dilution). Note that Fura-2 AM is photosensitive and should be kept in the dark and stored at 4  C. In addition to Fura-2 AM, Fura Red, Fluo3, or Fluo4 may be used with more readily available excitation/emission wavelengths as an alternative approach. 9. For calcium imaging, any imaging system capable of 340/380 nm paired excitation and 510 nm emission is sufficient. For fluorescent imaging, any imaging system capable of 488 nm excitation/525 nm emission (green fluorescence) and 535 nm excitation/620 nm emission (red fluorescence) is sufficient in order to differentiate with costaining. 10. Depending on experimental design, prepare glucose solution accordingly. 11. Blocking solution is prepared with 2% bovine serum albumin (BSA) and Triton X100 in 1 PBS.

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12. Starting near the base of the tail, open the abdomen using dissection scissors in a V-shape and pull the skin to expose abdominal and chest organs. Using gauze, gently push down on the diaphragm pulling back the liver to expose the common bile duct. 13. This is a critical step. 8 min is an average time frame. However, with different Collagenase lots as well as different mouse strains, this time may vary. It is important to be aware of this as the time may need to shorten or lengthen (approximately 5 min). 14. Approximately 2–4 h after isolation of islets it is advantageous to clean the islets. This can be done by transferring islets from one 10 mm petri dish with supplemented RPMI 1640 media to another 10 mm petri dish with supplemented RPMI 1640. Removing excess acinar tissue from the islets in the dish can reduce cell death and preserve islet function. Approximately 10 mL of supplemented RPMI media should be used per 300–350 islets. 15. To autoclave coverslips, use aluminum foil to make long rectangular packets that can hold 6–7 coverslips but no more than ¼ the width/diameter of the coverslip. Wrap the forceps in foil separately. 16. If needed, the coverslips may be broken into smaller pieces using an etching pencil and placed into a 6-well plate using the sterilized forceps, then coated with poly-D-lysine. To maintain the sterility of the poly-D-lysine solution, swab the stopper with an alcohol swab and use a 1 mL 25G TB syringe to draw off enough poly-D-lysine to completely cover the coverslips to the edge without overflowing off of the edge. Once coated, use caution to not scratch the coverslips as this can remove poly-Dlysine coat. 17. The larger the islet, the better for dissociation, and be sure to minimize the amount of medium that is collected with the islets when placed into the trypsin. Do only one trypsin tube at a time as islets should not be left in trypsin for prolonged periods of time. Rapid pipetting will help break islets using mechanical force while trypsin is used to help chemically. If after a few minutes of rapid pipetting you still see islets, continue for a few more minutes of rapid pipetting. 18. The easiest way to aspirate is to carefully tilt the tube and insert the aspiration line and be sure to aspirate as much of the supernatant as possible. 19. The amount of RPMI 1640 that the dissociated islets can be resuspended in can vary depending on the density of cells desired on the glass coverslips. Note that cells will concentrate

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in the center of the drop, so you may place suspension either as a single drop or as several depending on what the cells will be used for. 20. When starting the peristaltic pump, be sure to start the perfusion with dH2O in order to make sure the peristaltic pump is cycling properly. This ensures that the diamond bath imaging chamber is not leaking. Note, it is important that prior to beginning the calcium imaging of islets that the peristaltic pump is switched from dH2O to a glucose solution as dH2O alone will lyse the cells. Switch from dH2O to glucose solution 15–30 min prior to calcium imaging. 21. The first region of interest defined by the user should consist of a box of the background, completely clear of any cells or islets. This allows for a more accurate intensity of islets and cells by removing the background readings. 22. Following fixation and washing with 1 PBS, coverslips can be kept in 1 PBS at 4  C short term before continuing with staining procedure.

Acknowledgments Rachel T. Scarl and William J. Koch contributed equally to this work. References 1. Benninger RKP, Hodson DJ (2018) New understanding of β-cell heterogeneity and in situ islet function. Diabetes 67(4):537–547 2. Benninger RKP, Piston DW (2014) Cellular communication and heterogeneity in pancreatic islet insulin secretion dynamics. Trends Endocrinol Metab 25(8):399–406 3. Head WS, Orseth ML, Nunemaker CS, Satin LS, Piston DW, Benninger RKP (2012) Connexin-36 gap junctions regulate in vivo first- and second-phase insulin secretion dynamics and glucose tolerance in the conscious mouse. Diabetes 61(7):1700–1707 4. Caicedo A (2013) Paracrine and autocrine interactions in the human islet: more than meets the eye. Semin Cell Dev Biol 24 (1):11–21 5. Johnston NR, Mitchell RK, Haythorne E, Pessoa MP, Semplici F, Ferrer J, Piemonti L, Marchetti P, Bugliani M, Bosco D, Berishvili E, Duncanson P, Watkinson M, Broichhagen J, Trauner D, Rutter GA, Hodson DJ (2016) Beta cell hubs dictate pancreatic islet

responses to glucose. Cell Metab 224 (3):389–401 6. Lei C-L, Kellard JA, Hara M, Johnson JD, Rodriguez B, Briant LJB (2018) Beta-cell hubs maintain Ca2+ oscillations in human and mouse islet simulations. Islets 10(4):151–167 7. Marciniak A, Selck C, Friedrich B, Speier S (2013) Mouse pancreas tissue slice culture facilitates long-term studies of exocrine and endocrine cell physiology in situ. PLoS One 8 (11):e78706 8. Stozˇer A, Gosak M, Dolensˇek J, Perc M, Marhl M, Rupnik MS, Korosˇak D (2013) Functional connectivity in islets of Langerhans from mouse pancreas tissue slices. PLoS Comput Biol 9(2):e1002923 9. Nunemaker CS, Satin LS (2004) Comparison of metabolic oscillations from mouse pancreatic beta cells and islets. Endocrine 25 (1):61–67 10. Nunemaker CS, Dishinger JF, Dula SB, Wu R, Merrins MJ, Reid KR, Sherman A, Kennedy RT, Satin LS (2009) Glucose metabolism,

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islet architecture, and genetic homogeneity in imprinting of [Ca2+](i) and insulin rhythms in mouse islets. PLoS One 4(12):e8428 11. Scarl RT, Corbin KL, Vann NW, Smith HM, Satin LS, Sherman A, Nunemaker CS (2019) Intact pancreatic islets and dispersed beta-cells both generate intracellular calcium oscillations

but differ in their responsiveness to glucose. Cell Calcium 83:102081 12. Carter JD, Dula SB, Corbin KL, Wu R, Nunemaker CS (2009) A practical guide to rodent islet isolation and assessment. Biol Proced Online 11:3–31

Methods in Molecular Biology (2021) 2346: 165–172 DOI 10.1007/7651_2020_307 © Springer Science+Business Media New York 2020 Published online: 18 August 2020

Promoter Pull-Down Assay: A Biochemical Screen for DNA-Binding Proteins Ryan R. Chaparian and Julia C. van Kessel Abstract Transcription factors are ubiquitous proteins that associate with promoter DNA and regulate gene expression through a variety of mechanisms. Understanding transcriptional control mechanisms requires in-depth investigation of the binding of transcription factors to the promoters they regulate. There are many in vivo and in vitro methods for testing the binding of a known protein to a promoter, such as chromatin immunoprecipitation and electrophoretic mobility shift assays. However, for these experiments, one must have a protein candidate to test and is not able to identify unknown proteins bound to a particular promoter. Thus, the promoter pull-down assay was developed to fill this void. This method uses DNA as bait to capture proteins that bind to a specific promoter, such as transcription factors, from cellular lysates. Coupled with other experiments, the promoter pull-down assay vastly improves the repertoire of methods available for defining regulatory complexes that influence transcription. Keywords Promoter, Transcription factor, Transcription, Regulation, Protein identification, Vibrio, Quorum sensing

1

Introduction The transcription of a gene is dictated by the configuration of the upstream regulatory DNA called a promoter. Understanding transcriptional regulation in both eukaryotes and prokaryotes requires the identification/characterization of the regulatory proteins that bind the promoter of interest. In cases where a known protein is hypothesized to bind to a specific DNA region, there are several methods for assessing binding of that protein, including chromatin immunoprecipitation (ChIP) and various in vitro protein-DNA interaction assays (e.g., electrophoretic mobility shift assays, fluorescence anisotropy, biolayer interferometry, surface plasmon resonance, fluorescence resonance energy transfer, etc.) [1]. In instances where it is required to test where a particular protein binds in a genome, ChIP coupled with next-generation sequencing (ChIP-seq) is a proven and reliable method for the identification of such binding sites [2]. However, it can also be the case that one desires to test what unknown proteins bind to a specific promoter (or any genomic

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region). The promoter pull-down method takes an unbiased, biochemical approach to identify proteins bound to a defined promoter. Proteins are captured by flowing cellular lysate over promoter-bound beads. Positive hits that specifically bind to the target DNA are identified by comparison to a negative control DNA substrate. For example, in experiments probing for transcription factors that bind to a particular promoter, a negative DNA substrate could be DNA from within an open reading frame that is not expected to be bound by transcription initiation factors. This technique has been highly successful at assaying transcription factors that regulate quorum-sensing genes in Vibrio bacteria [3]. The experimental design of the pull-down technique allows for the capture of many proteins at once, negates the requirement of affinity tags/antibodies, and is relatively inexpensive. This method has been reported previously [4]; however, we have implemented several modifications to improve cell lysis efficiency and minimize the capture of proteins that bind to DNA non-specifically. Below, we describe the materials and methods for this assay. We note that the combination of the promoter pull-down assay, ChIP-seq, and in vitro assays is likely to yield highly informative and comprehensive snapshots of the DNA-binding proteins that occupy specific promoters under various conditions.

2

Materials All reagents should be prepared at room temperature, unless otherwise noted, using dH2O from an ultrapure water system (Millipore).

2.1 Probe Preparation

1. 50 -TEG-biotin-labeled primers (see Note 1). 2. 10 mM dNTPs. 3. 5 Phusion HF buffer (New England Biolabs). 4. Template DNA (genomic or plasmid). 5. Phusion polymerase (New England Biolabs). 6. Amicon Ultra centrifugal concentration unit (0.5 mL, Millipore).

2.2 Buffers and Reagents

1. TE: 0.5 M Tris–HCl pH 8.0, 1 mM EDTA pH 8.0. 2. THES buffer: 50 mM Tris–HCl pH 7.5, 10 mM EDTA, 20% sucrose, 140 mM NaCl, 1 protease inhibitors (see below). 3. 5 BS buffer: 50 mM HEPES pH 7.5, 25 mM CaCl2, 250 mM KCl, 60% glycerol. 4. BS/THES buffer: 44.3% THES buffer, 20% BS buffer.

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5. 2 B/W buffer: 10 mM Tris–HCl pH 7.5, 1 mM EDTA, 2 M NaCl. 6. 100 protease inhibitors mix: 0.07 mg/mL phosphoramidon (Santa Cruz), 1.67 mg/mL AEBSF (DOT Scientific), 0.07 mg/mL pepstatin A (DOT Scientific), 0.07 mg/mL E-64 (Gold Bio), 0.06 mg/mL bestatin (MP biomedicals/ Fisher). Protease inhibitors mix should be resuspended in DMSO. 7. NaCl elution buffers: 25 mM Tris–HCl pH 7.5, with varying concentrations of NaCl (100, 200, 300, 500, 1000 mM). 2.3

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1. 10 FastBreak cell lysis reagent (Promega). 2. Sonicator (Branson SFX 150 with microtip attachment).

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1. Dynabeads Scientific).

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2. DynaMag-2 magnetic stand (Thermo Fisher Scientific). 3. 1.5 mL microfuge tubes (Eppendorf). 4. Tube agitator (Thermo Fisher Scientific). 5. Pierce Streptavidin agarose resin (Thermo Fisher Scientific). 6. UltraPure salmon sperm DNA (Thermo Fisher Scientific). 7. Nuclease-free H2O. 2.5

SDS-PAGE Gel

1. Gel plates, combs, and gel chamber (standard systems). 2. SDS-PAGE buffers (standard Laemmli systems). 3. Plastic dish for gel staining. 4. Pierce silver stain kit (Thermo Fisher Scientific).

3

Methods All steps should be performed at room temperature unless otherwise noted. Care should be taken to avoid nuclease contamination, which will degrade the DNA probe and compromise the assay.

3.1

Cell Culture

1. Grow a 50 mL culture of the desired bacterial strain under the desired experimental conditions (see Note 2). 2. Record the optical density at 600 nm (OD600) per mL of culture. 3. Pellet the culture in a 50 mL conical tube by centrifugation at 1250  g for 10 min; remove and discard supernatant. 4. Pellet can be stored at 80  C for ~6 months or proceed to Subheading 3.2.

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3.2 Probe Preparation

1. Using biotinylated primers, prepare 10–12 PCR reactions (see Note 3). 2. Pool all reactions, and concentrate the DNA to 200 ng/μL using the Amicon Ultra centrifugal concentration unit (see Note 4). 3. If needed, dilute the probe to 200 ng/μL with nuclease-free ddH2O.

3.3 Dynabead Preparation

1. Add 100 μL of Dynabeads (1 mg) to a 1.5 mL microfuge tube. 2. Immobilize beads with a magnetic stand and remove storage solution (see Note 5). 3. Add 500 μL 2 B/W (binding/wash) buffer and incubate for 30 s. 4. Immobilize beads with magnetic stand and remove buffer. 5. Repeat steps 3 and 4 twice, for a total of three washes.

3.4 Probe Immobilization

1. Following the final wash from Subheading 3.3, add 200 μL of the 2 B/W buffer to the tube containing the washed Dynabeads. 2. Add 200 μL of 200 ng/μL concentrated DNA probe. 3. Incubate the bead and DNA probe mixture at room temperature with agitation for 20 min. 4. Immobilize the beads using the magnetic stand, and remove the supernatant (see Note 6). 5. Add 500 μL 2 B/W buffer and incubate for 30 s. 6. Wash the bead-DNA complex with 500 μL 0.5 M TE pH 8.0. 7. Immobilize the beads using the magnetic stand and remove the supernatant. 8. Repeat steps 6 and 7 for a total of three washes.

3.5 Bead Equilibration

1. Wash the DNA-bound beads with 500 μL of BS/THES buffer; incubate at room temperature for 1 min. 2. Immobilize the beads using the magnetic stand and remove the supernatant. 3. Repeat steps 1 and 2 for a total of two washes. 4. Add 500 μL of BS/THES buffer supplemented with 10 μg/ mL salmon sperm DNA. 5. Allow the probe-bound beads to incubate with the BS/THES buffer and salmon sperm DNA until the cell lysate has been prepared (Subheading 3.6).

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1. Resuspend the cell pellet with 1 FastBreak (diluted with BS/THES buffer) to 80–90 ODs/mL; incubate at 4  C with agitation for 30 min. 2. Move the cell lysate to a microfuge tube and chill on ice. 3. Sonicate the cell suspension for 15 s (at 30% amplitude) followed by a 1-min rest on ice (see Note 7). 4. Repeat step 3 for a total of five rounds of sonication.

3.7 Preclearing the Lysate

1. Pellet 1 mg of streptavidin-conjugated agarose beads via centrifugation at 700  g for 1 min (see Note 8). 2. Resuspend the beads in 500 μL BS/THES; incubate at room temperature for 1 min. 3. Pellet the beads via centrifugation at 700  g for 1 min. 4. Repeat steps 1 and 2 for a total of two washes. 5. Resuspend the immobilized beads with the sonicated lysate; incubate at room temperature for 30–60 min with agitation (see Note 9). 6. Move the precleared lysate to a clean microfuge tube.

3.8 Dynabead Incubation with Lysate

1. Immobilize the probe-bound Dynabeads from Subheading 3.5 using the magnetic stand, and remove the BS/THES buffer. 2. Resuspend the beads in 750 μL fresh BS/THES buffer, and add 25–100 μg salmon sperm DNA (see Note 10). 3. Add 200 μL of the precleared lysate to the beads. 4. Incubate at room temperature with agitation for 60 min. 5. Immobilize the beads using the magnetic stand and remove the supernatant. 6. Wash the beads with 500 μL fresh BS/THES buffer. 7. Immobilize the beads using the magnetic stand and remove the supernatant. 8. Repeat steps 6 and 7 for a total of five washes (see Note 11).

3.9

Elution

1. Resuspend the beads with 60 μL 100 mM NaCl elution buffer; incubate at room temperature for 1–2 min, with agitation. 2. Immobilize beads using magnetic stand; remove and save the supernatant for analysis. 3. Repeat steps 1–3 using 200, 300, 500, and 1000 mM NaCl elution buffers. 4. To completely elute all proteins and the DNA probe itself, resuspend the beads in nuclease-free ddH2O, and incubate at 70  C for 5 min (see Note 12).

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5. Immobilize the beads using the magnetic stand; remove and save the supernatant for analysis. 3.10 Analysis of Eluted Proteins

1. Pour an SDS-PAGE gel (see Note 13). 2. Load 10–20 μL of each elution sample (100–1000 mM NaCl elution samples and the 70  C elution sample). 3. Run the gel at ~13–16 V/cm (approximately 160 V for a standard mini gel apparatus). 4. Visualize proteins using a silver staining kit that is compatible with mass spectrometry. 5. Excise protein bands of interest, destain, and proceed to protein identification using mass spectrometry (see Note 14).

4

Notes 1. Primers can be ordered with a 50 -TEG-biotin modification from Integrated DNA Technologies. 2. This protocol was optimized for prokaryotes but can be modified for use with eukaryotic organisms. Escherichia coli cells contain ~0.2 pg of total protein per cell, whereas HeLa cells contain ~300 pg per cell. Thus, HeLa cells have approximately 1500 more protein per cell. This protocol uses ~4  1010 bacterial cells which equates to ~2.7  107 mammalian cells. However, the number of cells will depend on the desired OD600/conditions. 3. Multiple micrograms of biotinylated probe are required for this assay; thus many PCRs are necessary. PCRs contain 1 HF Phusion polymerase buffer (New England Biolabs), 0.2 mM dNTPs, 0.25 μM each primer, ~1 unit of Phusion polymerase (New England Biolabs). However, most standard PCR protocols are compatible with probe synthesis. Typically, 10–12 PCRs (50 μL reactions) are sufficient. 4. The Amicon Ultra centrifugal concentration units retain DNA between 137 and 1150 base pairs in size; primers are removed during concentration. 5. Complete immobilization of the beads requires approximately 1–2 min; however, this process is dependent on buffer viscosity and magnet strength. Bead immobilization can be monitored by proxy via buffer opacity – when the beads are fully immobilized, the buffer will return to completely transparent. 6. The DNA concentration can be measured (and should be dramatically lower) in the removed supernatant to ensure that the biotinylated DNA probe is efficiently binding the beads.

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7. Sonication is essential for disrupting genomic DNA and lipid membranes in the cellular lysate which would impair the efficiency of the pull-down. When sonicating, ensure that the tip is fully submerged in the lysate before applying power. Avoid generating bubbles with the sonicator, as heat dissipation becomes severely reduced and can result in decreased solubility of cellular proteins. 8. This step serves to deplete the lysate of “sticky” proteins and proteins that interact with streptavidin. Removing these proteins dramatically improves the quality of the assay by minimizing the number of false-positive protein hits. While streptavidin-conjugated agarose beads are the best choice for preclearing, any agarose resin can be used. 9. The purpose of this step is to deplete the lysate of proteins that bind to the beads and streptavidin in a non-specific manner. The number of cells used may change depending on the experimental design; however the 1 mg of streptavidin-conjugated agarose beads used for preclearing the lysate should be sufficient to efficiently remove “sticky” proteins from most lysate preparations. 10. The salmon sperm DNA serves as a non-specific competitor to the DNA probe immobilized to the beads. Other competitor substitutes, such as poly[dI-dC], calf thymus DNA, or heparin, can be used in place of the salmon sperm DNA. However, the inclusion of some kind of competitor is critically important for minimizing the number of non-specific DNA binding proteins that are captured. 11. The supernatants from the wash steps can be saved and analyzed via SDS-PAGE to gauge wash efficacy. These samples can also determine whether positive control proteins (i.e., proteins that should bind the probe DNA) are enduring the wash steps. 12. The streptavidin-biotin interaction is unstable at 70  C, and thus this incubation allows complete elution of the DNA probe and all remaining proteins. 13. High percentage polyacrylamide gels are desirable to ensure all proteins, especially small proteins, are identified. Gels containing 15–20% polyacrylamide are suitable; however, 4–20% gradient gels are optimal. 14. In order to identify positive protein hits, a negative control experiment needs to be analyzed in parallel. A negative control pull-down uses a DNA probe in which transcription factors/ promoter-binding proteins presumably do not bind. We have had success using intragenic DNA as a negative control probe.

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Positive hits are proteins that are present and significantly enriched in the elution samples from the experimental probe compared to the control probe. To ensure consistency, design the experimental and control probes to be the same size/GC content. References 1. Dey B, Thukral S, Krishnan S, Chakrobarty M, Gupta S, Manghani C, Rani V (2012) DNA-protein interactions: methods for detection and analysis. Mol Cell Biochem 365 (1–2):279–299. https://doi.org/10.1007/ s11010-012-1269-z 2. Marinov GK (2018) A decade of ChIP-seq. Brief Funct Genomics 17(2):77–79. https://doi.org/ 10.1093/bfgp/ely012 3. Chaparian RR, Olney SG, Hustmyer CM, Rowe-Magnus DA, van Kessel JC (2016)

Integration host factor and LuxR synergistically bind DNA to coactivate quorum-sensing genes in Vibrio harveyi. Mol Microbiol 101 (5):823–840. https://doi.org/10.1111/mmi. 13425 4. Jutras BL, Verma A, Stevenson B (2012) Identification of novel DNA-binding proteins using DNA-affinity chromatography/pull down. Curr Protoc Microbiol. Chapter 1:Unit1F 1. Doi: https://doi.org/10.1002/ 9780471729259.mc01f01s24

Methods in Molecular Biology (2021) 2346: 173–182 DOI 10.1007/7651_2020_306 © Springer Science+Business Media New York 2020 Published online: 24 July 2020

Purification of the Vibrio Quorum-Sensing Transcription Factors LuxR, HapR, and SmcR Jane D. Newman and Julia C. van Kessel Abstract Quorum sensing is a cell density-dependent form of cellular communication among bacteria. This signaling process has been heavily studied in vibrios due to their diverse and complex phenotypes and relevance to human and aquaculture disease. Mechanistic studies of Vibrio quorum sensing have required optimization of protein purification techniques to examine the role of key proteins, such as the LuxR/HapR family of transcription factors that control quorum-sensing gene expression. Protein purification is the cornerstone of biochemistry, and it is crucial to consistently produce batches of protein that are pure, active, and concentrated to perform various assays. The methods described here are optimized for purification of the Vibrio master quorum-sensing regulators, LuxR (Vibrio harveyi), HapR (Vibrio cholerae), and SmcR (Vibrio vulnificus). We anticipate that these methods can be applied to other proteins in this family of transcription factors. Keywords Protein purification, Transcription factor, Affinity purification, Native purification, HapR, LuxR, SmcR, Vibrio

1

Introduction Quorum sensing (QS) is a cell density-dependent communication system that is widespread among bacteria. QS regulates important phenotypes associated with density-dependent behavior such as virulence factor secretion, biofilm formation, protease production, and motility. In Vibrio QS systems, gene expression is primarily regulated by a family of TetR-type transcription factors that include LuxR (Vibrio harveyi), SmcR (Vibrio vulnificus), HapR (Vibrio cholerae), OpaR (Vibrio parahaemolyticus), LitR (Vibrio fischeri), VanT (Vibrio anguillarum), ValR (Vibrio alginolyticus), and VcpR (Vibrio coralliilyticus) [1]. Our methods describe specific purification schemes for LuxR, SmcR, and HapR, but we hypothesize that these methods should be applicable to other Vibrio LuxR/HapRtype proteins due to the high sequence conservation in this family. Specifically, HapR and SmcR have similar quaternary structures [2, 3], and both proteins share high amino acid identity with LuxR (71% and 92%, respectively). We have found that these LuxR-type QS transcription factors function best without tags or

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with only small N-terminal tags (e.g., FLAG, 6x-His) and are non-functional with large N-terminal tags (e.g., GST; Chaparian and van Kessel, unpublished). Based on the domain organization of these proteins, C-terminal tags would likely interfere with protein dimerization [3]. To facilitate biochemical assays with these important transcription factors, it is necessary to produce pure, active, and concentrated protein preparations. Depending on the downstream assay, different purification schemes may be optimal to generate either native protein (untagged) or tagged protein preparations. Despite the sequence-level similarity between these Vibrio proteins, we have found that the optimal purification scheme for LuxR is different from the optimal purification of SmcR and HapR [1]. In our hands, native purification of LuxR protein is easily accomplished using a combination of a Heparin affinity column, a Q column for anion exchange, and size exclusion chromatography [4, 5], whereas purification of SmcR and HapR using the same methods is unreliable. Thus, we developed two purification schemes for SmcR: an affinity purification using an N-terminal 6xHis-tag on SmcR and Ni-NTA resin system [6] and an affinity purification technique using InteinMediated Purification with an Affinity Chitin-binding Tag (IMPACT) to generate an untagged preparation of SmcR. For HapR purification, the 6xHis-tag and Ni-NTA purification is also successful.

2

Materials All reagents should be prepared at room temperature, unless otherwise noted and using dH2O from an ultrapure water system (Millipore).

2.1 Cell Culture Reagents

1. Lysogeny Broth (LB) Media: 10 g tryptone, 5 g yeast extract, 10 g NaCl, (15 g agar for plates) in 1 L dH2O. Autoclave. (a) Prepare 100 mL of LB in a bottle. (b) Prepare 2 1 L of LB in either a 2 L Erlenmeyer flask or 2.8 L Fernbach flask.

2.2 Buffers and Reagents

1. Native running buffer: 50 mM Tris pH 8, 50 mM NaCl. 2. Native elution buffer: 50 mM Tris pH 8, 1 M NaCl. 3. Ni-NTA running buffer: 25 mM Tris pH 8, 500 mM NaCl. 4. Ni-NTA elution buffer: 25 mM Tris pH 8, 500 mM NaCl, 500 mM imidazole. 5. IMPACT running buffer: 25 mM Tris pH 8, 500 mM NaCl, 1 mM EDTA.

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6. IMPACT wash buffer: 25 mM Tris pH 8, 1 M NaCl, 1 mM EDTA. 7. IMPACT cleavage buffer: 30 mL IMPACT running buffer, 231 mg dithiothreitol (DTT). 8. 1 M IPTG. 2.3

Cell Lysis

1. 10 FastBreak cell lysis reagent (Promega) or 10 BugBuster cell lysis reagent (Millipore). 2. Microfluidizer or emulsifier. 3. Lysozyme enzyme. 4. DNaseI enzyme (New England Biosciences, NEB). 5. Phenylmethylsulfonyl (Sigma).

(PMSF)

serine

protease

inhibitor

6. 100 protease inhibitors mix: 0.07 mg/mL phosphoramidon (Santa Cruz), 1.67 mg/mL AEBSF (DOT Scientific), 0.07 mg/mL pepstatin A (DOT Scientific), 0.07 mg/mL E-64 (Gold Bio), 0.06 mg/mL bestatin (MP Biomedicals/ Fisher). This is resuspended in DMSO. 2.4 Protein Purification

¨ kta 1. Fast Protein Liquid Chromatography (FPLC) system (A Pure from GE Healthcare Life Sciences) (see Note 1). 2. Snakeskin dialysis tubing (Thermo Fisher Scientific). 3. 5 mL Heparin column (GE Healthcare Life Sciences). 4. 5 mL Q column (GE Healthcare Life Sciences). 5. 5 mL Ni-NTA column (GE Healthcare Life Sciences) or Ni-NTA resin for gravity column (NEB). 6. Chitin resin (NEB). 7. Disposable gravity columns (Pierce, 10 mL column and plastic funnel). 8. 120 mL HiLoad™ 16/600 Superdex™ 75 pg column (GE Healthcare Life Sciences). 9. Vivaspin Turbo 10–15 kDa MWCO centrifugal filters (Sartorius®).

2.5

SDS-PAGE Gel

1. Gel plates, gel box, heating block, power supply. 2. SDS-PAGE buffers (standard Laemmli systems). 3. Plastic dish for gel staining. 4. Coomassie staining and destaining solutions.

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2.6 Autoinduction Media (Optional)

1. 2  1 L of autoinduction media made of the following reagents. (a) ZY Media (20 g tryptone and 10 g yeast extract in 1860 mL dH2O). (b) 1 M MgSO4 sterile filtered. (c) 50 5052 Media: 25% glycerol w/v, 2.5% glucose w/v, 10% alpha-lactose w/v, autoclaved. (d) 20 NPS (2 M (NH4)2SO4, 1 M KH2PO4, 1 M Na2HPO4) with a final pH of ~6.75 and autoclaved, made as shown in the table below: H2O

800 mL

(NH4)2SO4

66 g

KH2PO4

136 g

Na2HPO4

142 g

Final volume

1L

(e) 1000 trace metals mix is used because the concentration of metals becomes the limiting reagent for cellular growth in this medium. The 1000 trace metals mix is made with the stock solutions listed in the following table, each added to 36 mL dH2O using the volumes in the first column. Note that the 0.1 M FeCl3 should be prepared in 1/100 volume concentrated HCl instead of dH2O. Autoclave the stock solutions of the individual metals prior to making the trace metals mix except for the 0.1 M FeCl3 (HCl should never be autoclaved).

Stock solutions

MW (g/mol)

To make stocks: g/x mLa

1000 Conc. (mM)

1 Conc. In media (final) (μM)

50

0.1 M FeCl3-6H2O

270.30

13.52 g/500 mL

50 Fe

50 Fe

2

1 M CaCl2

110.99

11.10 g/100 mL

20 Ca

20 Ca

1

1 M MnCl2-4H2O

197.91

9.90 g/50 mL

10 Mn

10 Mn

1

1 M ZnSO4-7H2O

287.56

14.38 g/50 mL

10 Zn

10 Zn

1

0.2 M CoCl2-6H2O

237.95

2.38 g/50 mL

2 Co

2 Co

2

0.1 M CuCl2-2H2O

170.486

1.70 g/100 mL

2 Cu

2 Cu

1

0.1 M NiCl2-6H2O

237.72

2.38 g/50 mL

2 Ni

2 Ni

Volume (mL)

(continued)

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Stock solutions

MW (g/mol)

To make stocks: g/x mLa

1000 Conc. (mM)

1 Conc. In media (final) (μM)

2

0.1 M Na2MoO4-5H2O

241.98

2.42 g/100 mL

2 Mo

2 Mo

2

0.1 M Na2SeO3-5H2O

263.03

2.63 g/100 mL

2 Se

2 Se

2

0.1 M H3BO3

61.83

0.618 g/100 mL

2 H3BO3

2 H3BO3

Volume (mL)

a

All stock solutions should be prepared in dH2O except for 0.1 M FeCl3, which should be prepared in 1/100 volume concentrated HCl

(f) ZYP-5052 Media: 928 mL ZY media, 1 mL of the 1 M MgSO4 stock, 1 mL of 1000 tract metals mix, 20 mL of 50 5052, 50 mL of 20 NPS. (g) Filter the ZYP-5052 media prior to storing or use immediately, adding antibiotics for selection as needed (see Note 2).

3

Methods All steps should be performed with cells and/or protein on ice and/or in 4  C cold room.

3.1 Protein Overexpression (See Note 2)

1. The desired plasmid for overexpression should be introduced into E. coli BL21 (DE3) cells by transformation. 2. Inoculate 2  10 mL LB media cultures with selective antibiotics from single colonies or glycerol stocks and grow shaking at 30  C (see Note 3). 3. Dilute cells 1:100 into autoclaved 1 L LB media in flasks (add 10 mL from overnight cultures to each 1 L LB media). Grow shaking at 30  C until culture reaches an OD600 (optical density at 600 nm) between 0.4 and 0.6. 4. Induce protein expression with IPTG at a final concentration of 1 mM (1 mL of 1 M IPTG stock per 1 L culture) and grow shaking for 4 h at 30  C (see Note 4). 5. Centrifuge cells at max speed for at least 15 min, discard the supernatant, and freeze pellets at 80  C.

3.2

Cell Lysis

1. Resuspend the cell pellet in 50 mL running buffer. Use a stir bar to help resuspend the cell pellet for no longer than 20 min at room temperature. The buffer contents depend on which protein purification is to be used (see Subheading 2.2). 2. Add 200 μL100 mM PMSF, 50 μL10 mg/mL lysozyme, 200 μL 1 protease inhibitor, and 200 μL DNaseI to resuspended cell pellet, and homogenize sample thoroughly using a

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glass homogenizer or by serological pipetting to thoroughly disrupt cell clumps (must use homogenizer for mechanical lysis). 3. Lyse cells using chemical lysis (using 1 BugBuster reagent or 1 FastBreak reagent) or mechanical lysis (microfluidizer or emulsifier; see Note 5). 4. Centrifuge lysate at max speed for 30 min at 4  C to clarify lysate (see Note 6). 3.3 Protein Purification: Native Purification (See Note 7)

1. Equilibrate a 5 mL Heparin column using 5 column volumes (CV) of native running buffer using an FPLC. 2. Apply the protein lysate onto the Heparin column using either a sample pump, syringe, or Superloop. 3. Wash the Heparin column with 10 CV native running buffer. 4. Elute in 20 CV native elution buffer in a linear gradient ranging from 0 to 100% elution buffer (see Note 8). 5. Run 5 μL of elution samples on an SDS-PAGE gel to confirm presence of protein. 6. Pool peak UV fractions and place on ice. 7. Equilibrate a 5 mL Q column using 5 CV native running buffer. 8. Apply the pooled protein sample onto the Q column using either a sample pump, syringe, or Superloop. 9. Wash the Q column with 10 CV native running buffer. 10. Elute from the Q column with native running and elution buffers in a linear gradient ranging from 0 to 100% elution buffer (see Note 8). 11. Pool the protein fractions and use Size Exclusion Chromatography Subheading 3.6 if needed (see Note 9).

3.4 Protein Purification: Affinity Tag Option 1 – His-Tag (See Note 10)

1. Equilibrate a Ni-NTA column with 5 CV Ni-NTA running buffer. 2. Apply the protein lysate onto the Ni-NTA column using either a sample pump, syringe, or Superloop. 3. Wash the Ni-NTA column with 10 CV Ni-NTA running buffer. 4. Elute protein with a linear gradient over 20 CV of Ni-NTA elution buffer ranging from 0 to 100% elution buffer (see Note 8). 5. Run 5 μL of elution samples on an SDS-PAGE gel to confirm presence of protein. 6. Pool the protein fractions and use Size Exclusion Chromatography Subheading 3.6 if needed (see Note 9).

Purification of the Vibrio Quorum-Sensing Transcription Factors LuxR. . .

3.5 Protein Purification: Affinity Tag Option 2 – IMPACT (See Note 11)

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1. Remove 20 mL chitin slurry (equivalent to 10 mL chitin resin), and add to a disposable gravity column. Wash with 3 CV dH2O. 2. Equilibrate column with 10 CV IMPACT running buffer. 3. Apply the protein lysate to the chitin column. 4. Wash the chitin resin with 10 CV IMPACT running buffer. 5. Wash the chitin resin with 10 CV IMPACT wash buffer. 6. Incubate the chitin resin with 30 mL IMPACT cleavage buffer for 4 h to promote cleavage of the intein tag. 7. Elute protein with 2 CV IMPACT running buffer. 8. Dialyze sample for 2 h in gel filtration buffer prior to being applied to size exclusion chromatography, if needed (see Note 12). 9. Use Size Exclusion Chromatography Subheading 3.6 if needed (see Notes 9 and 13).

3.6 Protein Purification: Size Exclusion Chromatography (SEC) (See Note 13)

1. Concentrate protein prior to applying sample to size exclusion column using Sartorius® Vivaspin Turbo centrifugal filters with a 10 or 15 kDa MWCO (see Note 14). 2. Centrifuge at 4  C at 4000  g for 10 min intervals. 3. Gently pipet sample up and down between each spin, and continue centrifuging until volume of protein remaining is between 2 and 5 mL (see Note 15). 4. Manually inject 5 mL of gel filtration buffer, filtered and chilled. 5. Manually inject sample into sample port. 6. Run protein over the HiLoad™ 16/600 Superdex™ 75 pg column with the following protocol: 1.5 CV dH2O, 1.5 CV gel filtration buffer, sample application, and 1.5 CV gel filtration buffer for an isocratic elution (see Note 16). 7. Fractions collected from the elution are concentrated to desired concentration in the Sartorius® Vivaspin Turbo 10 kDa MWCO concentrators (see Notes 9 and 17).

4

Notes 1. Any pump-driven liquid chromatography system or gravity column can be used for the protein purifications described here. Always clean and store associated columns according to manufacturer’s instructions. 2. Autoinduction can be performed rather than an IPTG induction. This requires the reagents listed above in Subheading 2.6

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and the following method. Note that if the antibiotic used for selection is kanamycin, then omit NPS from the autoinduction media. Overnight cultures should be made using 10 mL LB media grown shaking overnight at 30  C in E. coli BL21(DE3). The next day the 10 mL culture should be added to 1 L of autoinduction media, and this is grown shaking 16–18 h at 18  C. The cells are then collected by centrifugation and the spent medium is discarded. Save the pellet at 80  C for up to 6 months. 3. Cells should be grown shaking at 30  C to keep proteins at optimal Vibrio growth temperature despite performing the overexpression in E. coli. 4. Overexpression of these proteins has the highest yield when using a T7 expression system. We recommend the pET28b vector in an E. coli BL21(DE3) strain. In a standard purification, 2 L of cells will yield >10 mg of LuxR/HapR/SmcR proteins. While IPTG induction is routinely used for these proteins, autoinduction has also been performed successfully. 5. Chemical lysis is sufficient for most biochemical assays, but mechanical lysis techniques are preferred for biophysical studies (e.g., crystallography). It is important to note that most machines that perform mechanical lysis should only have thoroughly homogenized cells and/or lysate applied, with no large cellular debris present. Sonication is not suggested as a method of lysis here because it is inefficient for the quantity of cells and can produce heat, which can denature heat-sensitive proteins. We recommend an Avestin EmulsiFlex-C3 emulsifier. 6. Pellet insoluble proteins and other cellular debris because the proteins of interest are soluble. Two rounds of centrifugation may be required to remove insoluble material. Also, the FPLC does not tolerate air bubbles in samples very well, especially if the air sensor function is being utilized for sample application. After centrifuging, filtering lysate samples with 0.45 μm filter before applying the lysate to the FPLC is a reasonable preparation step to avoid air bubble issues. 7. For biochemical studies in which a tag would interfere, a native purification is required. This protocol works for LuxR, but thus far has not worked for purifying SmcR or HapR. We recommend the IMPACT system for untagged SmcR protein. 8. Typically, the peak UV fractions elute at ~35% elution buffer. Approximately 2 mL of eluent is collected per tube during fractionator collection, but this could be reduced to 1 mL. 9. Protein intended for sensitive assays (e.g., crystal trays) should immediately be used or frozen in 80  C. Alternatively, for protein preparations intended for biochemical assays, add

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glycerol to a final concentration of 10%, flash-freeze in liquid nitrogen, and store at 80  C. 10. His-tagged protein purification is compatible with all LuxRtype proteins we have tested: LuxR, SmcR, and HapR. This can be performed using either an FPLC or by standard gravity flow with columns packed with 5 mL Ni-NTA resin. 11. The IMPACT purification has primarily been used to purify SmcR. The Intein tag is self-cleaving, and therefore the resulting purified SmcR can be used in assays that require no tag. We recommend using the pTXB1 plasmid that produces a C-terminal tag, but the tag is cleaved prior to any biochemical or structural analysis, which ablates any observable complications. The following purification method described is for a gravity flow purification that does not utilize FPLC. However, this can be successfully done used with FPLC as well by using hand-packed chitin columns. The chitin resin can be reused multiple times if properly cleaned and stored according to manufacturer’s instructions. 12. This dialysis step is important to decrease the concentration of DTT so that the protein does not crash out in gel filtration buffer upon being concentrated. 13. Depending on the purity required for planned experiments, either SEC or dialysis was performed as a final step. For the highest purity samples, SEC is required following either affinity or native purification. 14. Sartorius® Vivaspin Turbo centrifugal concentrators are optimal, with the least amount of protein adhering to the filter or crashing out of solution. 15. For the tightest resulting peak from SEC necessary for crystallography, concentrate to the smallest volume possible without protein crashing out of solution. It is challenging to concentrate below 1 mL, so we recommend stopping at this point. If small white flakes are observed, the protein has crashed out of solution, thereby yielding an unusable protein preparation. 16. If using an FPLC, the elution collection should be set up to collect anything above a threshold UV value of 5 mAU because the protein will have a much higher UV trace. Normally the samples come off in tubes 8–12 (2 mL fractions). For crystallography, the shoulders of the UV peak should not be used; save only the central peak samples. 17. Protein samples intended for crystallography studies should be concentrated to at least 5 mg/mL. We routinely obtain final preparations of ~2 mL at approximately 10–20 mg/mL of LuxR or approximately 4–8 mg/mL of SmcR and HapR.

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This amount of protein is only obtained when each step (overexpression, lysis, and purification) is performed optimally. These samples should be immediately used to set up a crystal tray, and remaining protein can be frozen at 80  C. References 1. Ball AS, Chaparian RR, van Kessel JC (2017) Quorum sensing gene regulation by LuxR/ HapR master regulators in vibrios. J Bacteriol 199:JB.00105-17. https://doi.org/10.1128/ JB.00105-17 2. Kim Y, Kim BS, Park YJ et al (2010) Crystal structure of SmcR, a quorum-sensing master regulator of Vibrio vulnificus, provides insight into its regulation of transcription. J Biol Chem 285:14020–14030. https://doi.org/10.1074/ jbc.M109.100248 3. De Silva RS, Kovacikova G, Lin W et al (2007) Crystal structure of the Vibrio cholerae quorumsensing regulatory protein HapR. J Bacteriol 189:5683–5691. https://doi.org/10.1128/JB. 01807-06 4. Van Kessel JC, Rutherford ST, Shao Y et al (2013) Individual and combined roles of the

master regulators apha and luxr in control of the Vibrio harveyi quorum-sensing regulon. J Bacteriol 195:436–443. https://doi.org/10. 1128/JB.01998-12 5. Chaparian RR, Olney SG, Hustmyer CM et al (2016) Integration host factor and LuxR synergistically bind DNA to coactivate quorumsensing genes in Vibrio harveyi. Mol Microbiol 101:823–840. https://doi.org/10.1111/mmi. 13425 6. Ball AS, van Kessel JC (2019) The master quorum sensing regulators LuxR/HapR directly interact with the alpha subunit of RNA polymerase to drive transcription activation in Vibrio harveyi and Vibrio cholerae. Mol Microbiol 111 (5):1317–1334. https://doi.org/10.1111/ mmi.14223

Methods in Molecular Biology (2021) 2346: 183–190 DOI 10.1007/7651_2020_305 © Springer Science+Business Media New York 2020 Published online: 18 August 2020

Preserving Cytonemes for Immunocytochemistry of Cultured Adherent Cells Sally Rogers and Steffen Scholpp Abstract Cytonemes are specialized signalling filopodia that have a role in development and cellular differentiation. However, they are not well preserved by standard fixation techniques to study protein localization and interactions. A recent methodological advance has yielded improvements in cytoneme preservation using glutaraldehyde fixation and sodium borohydride treatment to reduce background. We herein describe a safer method for effective blocking using glycine following glutaraldehyde fixation of cytonemes on cultured adherent cells and demonstrate its effectiveness in immunocytochemistry. Keywords Cytoneme, Filopodia, Fixation, Antibody, Staining, Confocal, Microscopy, Signaling

1

Introduction Cytonemes are thin, cellular projections that are specialized for exchange of signalling proteins between cells (reviewed in [1]). Standard fixation methods used for immunocytochemistry (e.g. using 4% paraformaldehyde) do not preserve cytoneme structures well. Therefore, there has been a reliance on overexpression of fluorescent recombinant proteins to determine interactions and localization of proteins suspected to be associated with signalling filopodia. A method for fixing cytonemes using 0.5% glutaraldehyde has recently been published [2]. This method uses sodium borohydride to remove the free aldehydes produced by the glutaraldehyde fixation that, if not dealt with, can result in non-specific antibody binding. Sodium borohydride reduces the aldehydes that remain in the sample, so that they are no longer reactive, preventing them from covalently crosslinking to antibodies. However, sodium borohydride is toxic, flammable and corrosive and causes a health hazard. It releases gas upon solution preparation and must be prepared fresh (within 10–15 min before use) to be effective. Taken together, it is desirable to use an effective, yet safer and easier, substitute for blocking prior to immunocytochemistry following glutaraldehyde fixation. Glycine, as a primary amine, reacts with any

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free aldehydes in much the same way as a protein would, neutralizing any remaining aldehyde groups. We have therefore developed a method to use glycine to block non-specific antibody binding following fixation of cytonemes with glutaraldehyde. Here, we describe this technique from plating out cultured adherent cells through fixation, blocking and subsequent antibody staining and demonstrate its effectiveness.

2

Materials Prepare all solutions using ultrapure water (prepared by purifying deionized water, to attain a sensitivity of 18 MΩ cm at 25  C) and analytical grade reagents. Prepare and store all reagents at room temperature (unless indicated otherwise). Diligently follow all waste disposal regulations when disposing of waste materials. We do not add sodium azide to reagents.

2.1

Cell Culture

1. Sterile coverslips: These should be appropriate for the subsequent microscopy. There are several ways to sterilize coverslips. Place coverslips into a glass beaker and cover with foil. Autoclave, and follow with a drying cycle. Alternatively, coverslips can be placed in 70% ethanol in a Petri dish for 15 min. During this time, place a paper towel in a laminar flow hood, and spray liberally until soaked with 70% ethanol. Place the coverslips onto the wet paper towel in the hood, and allow to dry (see Note 1). 2. Six-well plates (see Note 2). 3. Adherent cells. 4. Growth media as required by cell type, containing 10% FBS. 5. Trypsin-EDTA (0.05%). 6. 1 PBS.

2.2 Fixation, Permeabilization and Block

1. Sorensen’s phosphate buffer: 0.133 M Na2HPO4 and 0.133 M KH2PO4 (pH 7.4). Prepare 0.133 M dibasic Na2HPO4 (anhydrous) by adding 9.44 g to 500 ml dH2O. Prepare 0.133 M monobasic KH2PO4 by adding 9.05 g to 500 ml dH2O. To make pH 7.4 Sorensen’s phosphate buffer, mix 80.4 ml of 0.133 M Na2HPO4 with 19.6 ml of KH2PO4 (see Note 3). 2. MEM-fix: 0.1 M Sorensen’s phosphate buffer (pH 7.4), 4% formaldehyde and 0.5% glutaraldehyde (see Note 4). To prepare 30 ml of MEM-fix (see Note 5), add 26.7 ml Sorensen’s phosphate buffer pH 7.4 to a 50 ml tube. Add 3 ml of 38% formaldehyde and 300 μl of 50% glutaraldehyde. Prepare the day before use and store at 4  C.

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3. Permeabilization and blocking solution: 0.1 M Sorensen’s phosphate buffer (pH 7.4), 0.1% Triton X-100, 5% serum (species depending on the secondary antibody), and 0.2 M glycine (see Notes 6 and 7). To prepare 15 ml (see Note 8), add 13 ml of Sorensen’s phosphate buffer pH 7.4 to a tube (see Note 9). Add 150 μl 10% Triton-X 100, 1.5 ml of 2 M glycine and 750 μl of serum. Mix by inverting several times gently, and store at room temperature if prepared immediately before use, or at 4  C. 2.3 Immunocytochemistry

1. Incubation buffer: 1 dPBS, 5% goat serum, 0.1% Tween-20. To prepare 1.5 ml, aliquot 1.4 ml of 1 dPBS, and then add 75 μl of goat serum and 15 μl of 10% Tween-20. Mix by gently pipetting up and down. Prepare on day of use. 2. Tween wash buffer: 1 dPBS, 0.05% Tween-20. Add 500 μl of 10% Tween-20 to 100 ml dPBS. Invert several times gently to mix, and store at room temperature. 3. Antibody incubation chamber (see Note 10 and Fig. 2).

2.4 Mounting Coverslips

1. Slides suitable for microscopy. 2. Mountant: We use Invitrogen™ ProLong™ Diamond Antifade Mountant for its low background and anti-bleaching properties. 3. Nail varnish.

3

Methods Carry out all procedures at room temperature unless specified otherwise. Cell culture techniques should be performed in a dedicated laminar flow hood using sterile technique.

3.1

Cell Culture

1. For most experiments, we start with a 75 cm2 flask. Dissociate adherent cells from culture by removing the growth media, rinsing once in 10 ml of prewarmed 1 PBS and then incubating for 5–15 min in 3 ml of prewarmed Trypsin to cover the cells. Check cells every 5 min to identify when they have detached from the culture plastic. Add 10 ml of growth media containing 10% FBS to neutralize the trypsin and resuspend the cells. Collect to a sterile tube and count cells. Aliquot sufficient cells for your experiment, centrifuge at 300  g for 10 min, aspirate supernatant, and resuspend the cell pellet in a suitable volume of sterile growth media. 2. Using sterile tweezers, place a sterile coverslip into a well on a sterile six-well plate. Prepare enough wells to meet your experimental conditions (see Note 11).

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3. Aliquot the required number of cells (usually 1  105 to 5  105 cells; see Note 12) into the well in 2 ml final volume of growth media (see Note 13). 4. Incubate cells at 37  C, 5% CO2 for 24 h to allow cells to adhere fully to the coverslips. View cells to check confluency and for expression of any recombinant proteins before proceeding to the next step. 3.2 Fixation, Permeabilization and Blocking

1. From this point on, cells need to be handled very carefully to prevent breakage of cytonemes. Do not aspirate; instead buffers should be gently transferred using a Pasteur pipette. Incubations should be performed without rocking or agitation. 2. Remove culture plates from the incubator. Remove culture media. Rinse cells gently, but quickly, three times in prewarmed dPBS, discarding the wash each time (see Note 14). 3. Immediately add 2 ml of cold MEM-fix to the cells, and incubate for 7 min at 4  C. 4. If the cells are expressing fluorescent recombinant proteins, the following steps should be performed in low light conditions and incubated in the dark to avoid bleaching. 5. Wash 3 5 min in 1 Sorensen’s phosphate buffer pH 7.4 to remove the MEM-fix (see Note 15). 6. Permeabilize the cells by adding 1 ml of permeabilization and blocking buffer. Incubate for 1 h at room temperature (see Note 16).

3.3

Antibody Binding

1. Wash the cells on the coverslips 3 5 min in dPBS. Leave the last wash on the cells. 2. During the washes prepare the antibody incubation equipment (see Fig. 2). Each coverslip will require 30–50 μl of primary antibody solution depending on the size of the coverslip. Damp paper towel is included in the incubation chamber to prevent the coverslips from drying out. Place the required amount of incubation buffer into a sterile 1.5 ml tube on ice; then add the amount of antibody required for the optimum dilution (see Note 17). Pipette 30–50 μl of primary antibody solution onto the Parafilm in the incubation chamber (see Note 18). Ensure that the coverslips can be identified by labelling either the side of the chamber or noting the position of the coverslips. 3. Following the PBS washes, the coverslips should be placed onto the antibody solution in the chamber. Using tweezers and a 100 μl pipette tip, gently lift the coverslip from the well. Dab the edge of the coverslip onto some paper towel to remove excess liquid. Then place the coverslip, cell side down,

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onto the drop of antibody solution in the chamber. Gently use the pipette tip to squeeze out any air bubbles under the coverslip. Repeat for all of the coverslips. 4. Cover the incubation chamber (see Note 19) and incubate at 4  C overnight. 5. The following morning, prepare and label sufficient clean six-well plates for your coverslips. Gently lift the coverslips using tweezers, and place cell side up, into the well of the plate (see Note 20). Rinse 1 in Tween wash buffer; then wash 3 5 min in Tween wash buffer, followed by 2 5 min in dPBS. 6. During the washes, prepare the incubation chamber for the secondary antibody, by discarding the previous Parafilm and replacing with new. Prepare the appropriate dilution of secondary antibody in incubation buffer (see Note 21). It is possible to add further dyes at this stage, for example, Phalloidin at the recommended concentrations. Aliquot 30–50 μl of secondary antibody solution to the incubation chamber as before. 7. Following the washes, use tweezers and a pipette tip to lift the coverslips from the six-well plate, dab off excess liquid, and place cell side down onto the antibody solution. Take care to note the location or identity of the coverslips. Incubate for 1 h at room temperature in the dark. Remove the mountant from freezer storage, and allow to warm up to room temperature during the incubation. 8. During the incubation, prepare the slides. Wash thoroughly using water and white tissue, and leave to dry. Label clearly with a permanent marker, and lay out on paper towel (see Note 22). 3.4

Mounting

1. Following the incubation, repeat the washes in step 5 above. Remove the final PBS wash, and gently add 2 ml of water. Immediately remove the coverslip from the well using tweezers, drain off excess liquid onto a paper towel, and lay the coverslip cell side up next to its pre-labelled slide. Allow to air-dry for 5–10 min. 2. Place a drop of mountant to the centre of each slide, taking care to avoid bubbles. Carefully lift the coverslip using tweezers, and place cell side gently down onto the drop of mountant. Once all the coverslips are mounted, cover, and allow to dry for 24 h (or as according to specific manufacturer’s instructions). For semihard mountants, seal the edges of the coverslip with clear nail varnish, and store at 4  C in the dark. See Fig. 1 for final image.

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Fig. 1 Confocal microscope image of adherent gastric cancer cells prepared using the method described. Cells were incubated with primary antibody specific for a Wnt ligand, detected with a species-appropriate secondary antibody conjugated to AF405 (shown in blue) and Phalloidin FITC (shown in green). The scale bar in bottom right indicates 10 μm

4

Notes 1. It may be necessary to coat the coverslips with poly-D-lysine in addition dependent on cell line. 2. Six-well plates allow easy handling of the coverslips, but other formats could be used if more suitable to individual experiments. 3. These volumes should give the correct pH, but it is advisable to check. 4. The concentration of glutaraldyde can be lower to optimize for different antibodies. We have used 0.2% glutaraldehyde to fix the cytonemes and improve antibody binding. 5. This volume is sufficient for fixing 12 coverslips in 6-well plates. 6. The concentration of glycine can be increased to 0.3 M if you experience high background. 7. Other permeabilization agents may be substituted, depending on cell type and antibody, for example, 100 μM digitonin or 0.5% saponin. 8. This volume is sufficient for permeabilizing and blocking 12 coverslips in 6-well plates. 9. We have substituted 1 dPBS at this step with equally good results. 10. We prepare an incubation chamber using recycled, clean tip boxes as follows (Fig. 2). Add a layer of paper towel to the

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Fig. 2 Assembly of an antibody incubation chamber using a repurposed tip box. The antibody solution is placed on top of the Parafilm, and the coverslip placed cell face down on top of the antibody. The incubation chamber is kept humid with the addition of a layer of damp tissue paper or blotting paper underneath the Parafilm. The chamber can be covered in foil for light-sensitive antibodies

bottom of an empty tip box, and soak with water. Pour off any excess. Place a layer of Parafilm® cut to size on top of the wet paper towel. 30–50 μl of antibody-containing incubation buffer can be aliquoted onto the Parafilm, allowing the coverslips to be placed, cell side down, on top of the drop. 11. For the first few times of performing this technique, it is worth preparing at least two coverslips for each experimental condition in case of breakage. 12. Cell number will depend on your cell type and confluency required at staining. 13. It may be desirable to transfect cells before plating onto coverslips, for example, with fluorescent membrane markers or tagged proteins of interest for co-localization studies. In this case, transfect cells as usual, then dissociate, count, and plate the cells onto coverslips 24–48 h post-transfection. We have had success with GFP, AF488 and glycophosphatidylinositol (GPI)-anchored, membrane-bound mCherry (mem-mCherry) transfections using this approach. 14. We have washed just once in PBS to minimize retraction of cytonemes and still achieved excellent results. 15. It is possible to use 1 dPBS at this step but needs confirmation for each cell type/antibody combination. 16. Incubation time can be optimized for each cell type and antibody combination. Shorter incubation times may result in better epitope survival. 17. The optimum dilution will need to be determined for each antibody; in general, we start with 1/50 to 1/100 dilution. 18. We use a grid pattern, fitting 2  3 coverslips in each chamber. 19. We use the lid of the pipette box covered in foil. 20. If you drop the coverslips, it may be possible to identify the cell side of the coverslip by gently scraping the cells at one edge. If

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you break the coverslip, either use the backup, or it may be possible to continue depending on the size of the fragments. 21. Each combination will need optimizing; we start with 1/10,000. 22. It is helpful at this stage to lay out your slides in the same pattern as your coverslips are organized in your plates to help with the transfer, particularly if you have a large number of coverslips.

Acknowledgements We would like to thank Daniel Routledge for providing comments on the manuscript. This project was supported by the Medical Research Council (MRC) research grant MR/S007970/1 (awarded to SS). References 1. Zhang C, Scholpp S (2019) Cytonemes in development. Curr Opin Genet Dev 57:25–30 2. Bodeen W, Marada S, Truong A et al (2017) A fixation method to preserve cultured cell

cytonemes facilitates mechanistic interrogation of morphogen transport. Development 144 (19):3612–3624

Methods in Molecular Biology (2021) 2346: 191–206 DOI 10.1007/7651_2020_330 © Springer Science+Business Media New York 2020 Published online: 29 September 2020

Fluorescent Labeling of Connexin with As Complex and X-Y Coordinate Registration of Target Single Cells Based on a Triangle Standard Chip for the Image Analysis of Gap Junctional Communication Mikako Saito Abstract Gap junction (GJ) research has entered a new stage focusing the concerted dynamic behavior of multiple isoforms of connexin (Cx) in the cell membrane, cytosolic vesicles, and space between them. To proceed with this research, imaging technologies are important. Here we describe two novel protocols for this purpose. At first, the adoption of a small motif of Cys-Cys-X-X-Cys-Cys as a visualization tag is described. An As complex, FlAsH, can bind to this tetra-Cys (TC) tag to form a fluorescent conjugate. Its introduction into the C-terminal of Cx43 is demonstrated. Next, a novel triangle chip for the accurate x-y registration is described. Target single cells of HeLa marked with a fluorescent dye can be easily recognized by electron microscopy based on this chip. Key words Connexin, Tetra-Cys tag, Target single cells, Triangle standard chip

1

Introduction Gap junctional communication is well understood to be relevant to various growth stages, metabolic conditions, and diseases by facilitating direct intercellular movement of small molecules [1, 2]. A gap junction (GJ) comprises six connexin (Cx) molecules. There are 21 Cx isoforms in humans and 20 isoforms in mice [3]. Cx proteins are synthesized in the endoplasmic reticulum and transported to the Golgi apparatus [4]. During this transportation, a Cx molecule is folded into three-dimensional structure and then oligomerized into a hexameric hemichannel, a connexon. Connexons in the Golgi apparatus are packaged into vesicles and transported to the cell membrane. An individual connexon from one cell associates with a corresponding connexon on a neighboring cell to form a GJ channel. Usually multiple channels aggregate in the cell membrane to form GJ plaques [5]. Conversely, the process of decomposition into single Cx molecules is also understood to progress in the

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cytoplasm [6]. To date, such a general view of the dynamic behavior of Cx has been analyzed mostly by molecular biological methods with image data provided by fluorescent microscopy. Recently, special attention has been focused on the potential role of Cx as tumor suppressors [7]. The specific roles of several isoforms such as Cx43 and Cx26 have been well investigated because they are more ubiquitously expressed than other isoforms. Until now, however, it is still unclear whether they are suppressive [8, 9] or progressive [10, 11]. A breakthrough to find a solution to this problem should be given by the analysis of the spatiotemporal behavior of Cx and its associated clusters with as high resolution as possible. In this new research stage, it has been recognized to be important to pay special attention to such isoforms that might be formerly ignored because of low expression and short lifetime [12]. Consequently, we have focused our efforts on the improvement of live imaging applicable to fluorescent microscopy and on the zooming up its correlated images to the electron microscopy level. Green fluorescent protein (GFP) is a widely used probe, but its molecular size (ca. 25 kDa) is too large for a Cx molecule. The fusion of GFP to Cx might disturb the intrinsic function of Cx. Here we have adopted a small motif of Cys-Cys-X-X-Cys-Cys as a visualization tag [13] and introduced it into the C-terminal of Cx43 [14]. An As complex, FlAsH, can bind to this tetra-Cys (TC) tag to form a fluorescent conjugate. Regarding the correlative x-y coordinate system, special devices were developed by pioneering works of Haraguchi et al. [15–17], and their system has been called correlative light-electron microscopy (CLEM). They developed special glass dishes with typically a size of 35 mm in diameter and location markers such as grid lines and/or letters on the bottom. A lattice pattern, for instance, of 500 μm  500 μm, is formed with 40–50 μm thick grid lines. Those dishes are now commercially available. When cells are observed with an inverted fluorescent microscope, however, it should happen to occur that target cells with a size of no greater than 15 μm are hidden behind a grid line. Our idea was to use a right triangle plastic chip and its engraved shape [18] to prevent such an obstruction of field of microscopic view. There is no need to draw position markers such as grid lines, alphabet letters, and numbers. Such an idea is based on the idea of the coordinate microchip in Suguwaculture system [19, 20]. Here we describe two protocols: one for the fluorescent imaging of Cx43 clusters using a TC tag and the other for the unobstructed field of view in microscopy. HeLa cells are used throughout because there is no Cx in wild HeLa cells and useful for the analysis of only the specific isomer(s) of interest by genetic introduction of those isoform(s) into HeLa cells.

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2 2.1

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Materials HeLa Cell Culture

1. Culture medium for HeLa cells (HeLaM): 43.5 mL of DMEM (Sigma), 5 mL of 10% FBS, 0.5 mL of (100) nonessential amino acids (Gibco), 0.5 mL of 200 mM L-glutamine solution (Gibco), and 0.5 mL of 1 mM sodium pyruvate solution per 50 mL. 2. HeLa cell line obtained from Bioresource Center, RIKEN (Tsukuba, Japan). Regarding cell freezing for storage and thawing for use, refer to Note 1. 3. Cell passage: Culture cells in HeLaM in culture dishes (10 cmϕ) at 37  C under 5% CO2. When the cells become 70–80% confluent after culture for 4–5 days, replace the medium by PBS to wash the cells twice. Add 0.3 mL of trypsin-EDTA solution (0.25% trypsin, 1 mM EDTA, Thermo Fisher Scientific) to the dish for the reaction at 37  C for 5 min under 5% CO2. Then add 3 mL of HeLaM to the dish to suppress the trypsin reaction. Homogenize the suspended cells gently by pipetting, and transfer in a 15 mL centrifugation tube. Centrifuge at 350  g at 4  C for 5 min to collect cells as the precipitate. Suspend cells in 2 mL of HeLaM, dispense them in dishes (10 cmϕ) at 3  105 cells/dish, and culture at 37  C under 5% CO2.

2.2

Transfection

1. Culture medium for Escherichia coli 5α: LB broth (Tryptone 1%w/v, yeast extract 0.5%, NaCl 1% (w/v)) or LB agar medium (LB broth, agar 1.5%w/v). 2. Restriction enzymes (AgeI, EcoRI) and T4 DNA ligase (Invitrogen). 3. pDsRed monomer C1 vector (Clontech) (see Note 2). 4. Selection marker (G418): Prepare a 900 μg/μL solution. 5. Solutions for E. coli cell lysis: Solution I: 1 M Tris-HCl (pH 8.0) 2.5 mL, 0.5 M EDTA 2 mL, 1 M glucose 5 mL, and sterile water 90.5 mL (total 100 mL). Solution II: 1 M NaOH 1 mL, 10% SDS 0.5 mL, sterile water 3.5 mL (total 5 mL). Solution III: 5 M potassium acetate 60 mL, glacial acetic acid 11.5 mL, sterile water 28.5 mL (total 100 mL). 6. Culture media used in transfection into E. coli: SOC medium: SOB medium 120 mL, 1 M glucose solution 2.4 mL (total 122.4 mL). SOB medium: Bacto Tryptone 2.4 g, Bacto Yeast Extract 0.6 g, NaCl 0.06 g, KCl 0.0023 g, water (total 120 mL); add 10 μL/mL of 1 M MgCl2 and 1 M MgSO4 after autoclave, before use.

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2.3 Fluorescent Dyes for Diffusion Marker

1. Dextran, Texas Red (DTR) (Invitrogen): M.W. 70 kDa, Ex/Em 595/615 nm. 2. Lucifer yellow (LY) (Sigma-Aldrich): M.W. 443 Da, Ex/Em 428/554 nm. 3. Adjust the concentrations of DTR and LY in the injection capillary at 1 mg/mL and 0.4 mg/mL, respectively. DTR and LY are impermeable and permeable markers to confirm the gap junctional molecular permeability of GJs composed of Cx43, because GJ cutoff size is estimated as 1000–1500.

2.4 Chelating Reagent for a TC Tag and Washing Buffer

1. FlAsH (Toronto Research Chemicals Inc., Toronto, Canada): Prepare a 500 nM solution.

2.5

1. A polystyrene plate with 0.3 mm thick.

Polystyrene Plate

2.6 Cell Fixation Reagents

2. 2,3-Dimercapto-1-propanol (BAL) wash buffer solution (Invitrogen Molecular Probes).

1. 1.5% OsO4 in 0.1 M phosphate buffer saline (PBS). 2. Ethanol solutions (30%, 50%, 90%, and 100%). 3. EPON 812 and its ethanol solutions (10%, 30%, 50%, 70%, and 90%).

3

Methods

3.1 Preparation of an Insert DNA: A Cx43-TC Chain

1. Extract RNA from mouse ES cells (see Note 3). 2. Prepare cDNA from RNA by reverse transcription (see Note 4). 3. Design the forward primer comprising a sequence targeting Cx43 and an AgeI recognition adaptor at the 50 terminal (Fig. 1). 4. Design the reverse primer comprising a sequence targeting Cx43, tetracysteine (TC) tag ( AEAAAREACCRECCARA) and an EcoRI recognition adaptor at the 50 terminal (Fig. 1).

Fig. 1 Schematic diagram of primer design for a Cx43-TC chain

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5. Prepare a PCR reaction solution: A mixture of GoTaq® Green Master Mix 5 μL, 10 pmol/μL forward primer 0.4 μL, 10 pmol/μL reverse primer 0.4 μL, cDNA 1 μL, and H2O 3.2 μL (total 10 μL). 6. Conduct PCR under the condition: Initial denature 94  C 3 min, 30 cycles [denature 94  C 1 min, annealing 58  C 1 min, extension 72  C 2 min], final extension 72  C 2 min. 7. Purify the PCR product: Transfer 100 μL of the PCR product to a 1.5 μL microtube. Add 100 μL of neutral phenol/chloroform (1:1) solution to the microtube, fully mix, and conduct centrifugation at 20,000  g at 4  C for 5 min. Transfer the supernatant to another microtube, add 10 μL of 3 M sodium acetate and 250 μL of 99% ethanol to the microtube, and keep it at 80  C for 15 min. Conduct centrifugation at 20,000  g at 4  C for 20 min, addition of 70% ethanol to the precipitate, and again centrifugation at 20,000  g at 4  C for 5 min. Finally, air dry the precipitate for 5 min, and suspend in 10 μL Tris-EDTA (TE) buffer solution (pH 8.0) to obtain the purified PCR product. 8. Treat the purified PCR product with AgeI: Prepare a mixture of the purified PCR product 10 μL, AgeI 1.67 μL, 10 M buffer (Takara) (see Note 5) 5 μL, sterile water 33.33 μL (total 50 μL) for reaction at 37  C for 3 h. Conduct addition of 125 μL of 100% ethanol and 5 μL of 3 M sodium acetate to the mixture, keeping it at 80  C for 15 min, centrifugation at 20,000  g at 4  C for 20 min, addition of 200 μL of 70% ethanol to the supernatant, and again centrifugation at 20,000  g at 4  C for 5 min, and finally, suspend the precipitate in 10 μL sterile water to obtain the AgeI-treated product. 9. Treat the AgeI-treated product with EcoRI: Prepare a mixture of 10 μL of the AgeI-treated product, EcoRI 0.67 μL, 10 H buffer (Takara) (see Note 5) 5 μL, sterile water 34.33 μL (total 50 μL) for reaction at 37  C for 3 h. 10. Electrophorese the EcoRI-treated product at 100 V for 30 min using 1% agarose/1 TAE gel and 1 TAE electrophoresis buffer. Immerse the gel in the EB staining solution for 15 min and then in the decolorization solution for 15 min. Cut out the gel containing target DNA sequence under UV illumination (see Note 6), and transfer in a 1.5 mL microtube. Add 10 μL of Membrane Binding Solution to the microtube per 10 mg of gel, and incubate at 55  C for 10 min to dissolve the gel pieces. Place the dissolved solution in a spin column set, incubate for 1 min at room temperature, centrifuge the column at 16,000  g for 1 min, discard the eluted solution, add 700 μL of Membrane Wash Solution to the column, and centrifuge the column again at 16,000  g for 1 min. Discard

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the eluted solution, add 500 μL of Membrane Wash Solution to the column, and centrifuge the column at 16,000  g for 5 min. Transfer the column content to a new 1.5 mL microtube, add 50 μL of elution buffer, incubate for 1 min at room temperature, and centrifuge at 16,000  g for 1 min to obtain the eluted solution as the purified Cx43-TC chain. 3.2 Isolation of a Plasmid Vector pDsRed Monomer C1

1. Spread E. coli transfected with pDsRed monomer C1 on LB plates, and culture at 37  C for 12–14 h. Pick up a single colony, inoculate it in LB medium containing 20 μg/mL kanamycin (Km), and culture at 37  C for 12–14 h with rotation at 170 rpm. 2. Collect E. coli cells by centrifugation at 630  g for 5 min. Add 100 μL of solution I to the precipitation and fully suspend cells. Add 200 μL of solution II for gentle stirring, and add 150 μL of solution III for full stirring. 3. Add 100 μL neutral phenol/chloroform (1:1) for full stirring, and centrifuge at 20,000  g at 4  C for 3 min. Transfer the supernatant to a microtube, add 500 μL isopropyl alcohol, centrifuge at 20,000  g at 4  C for 3 min, and discard the supernatant. Add 500 μL of 70% ethanol to the precipitate for full stirring, centrifuge at 13,000  g at 4  C for 5 min, and discard the supernatant. Air dry the precipitate at 37  C for 5 min, add 100 μL of TE (pH 8.0) for full stirring, and add 0.5 μL RNase (Nippon Gene) for incubation at 37  C for 30 min to obtain the crude sample of the plasmid vector. 4. Dispense the crude vector solution to microtubes, and add the same volume of 13% polyethylene glycol (PEG)/0.8 M NaCl for full mixing and then incubation in ice water for 1 h. Centrifuge at 20,000  g at 4  C for 20 min and discard the supernatant. Dissolve the precipitate with 200 μL TE, and add 200 μL neutral phenol/chloroform (1:1). Transfer to a new microtube, add 500 μL of 100% ethanol and 20 μL of 3 M sodium acetate, and incubate at 80  C for 15 min. Centrifuge at 20,000  g at 4  C for 10 min, discard the supernatant, wash the precipitate with 70% ethanol, and dissolve with 30 μL TE. Check the purity of the plasmid vector pDsRed monomer C1 from its absorbance.

3.3 Linearization of the Plasmid Vector

1. Treat pDsRed monomer C1 with AgeI and EcoRI: Prepare a mixture of 10 μL of pDsRed monomer C1, 1.67 μL of AgeI, 5 μL of 10 M buffer (see Note 5), and 33.33 μL of sterile water (total 50 μL) for incubation at 37  C for 30 min. Add 125 μL of 100% ethanol and 5 μL of 3 M sodium acetate for incubation at 80  C for 15 min. Centrifuge at 20,000  g at 4  C for 20 min to collect the supernatant, add 200 μL of 70% ethanol, and centrifuge at 20,000  g at 4  C for 5 min to

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discard the supernatant. To the precipitate, add 10 μL of sterile water, 0.67 μL EcoRI, 5 μL of 10 H buffer (see Note 5), and 34.33 μL of sterile water (total 50 μL) for incubation at 37  C for 30 min. 2. Electrophorese the EcoRI-treated product to obtain the purified vector DNA in the same procedure as Subheading 3.1, step 10. 3.4 Ligation of Cx43TC Chain and the Vector to Generate pCMV-Cx43-TC

1. Ligate the Cx43-TC chain with the linearized plasmid vector: Prepare a mixture of the purified Cx43-TC chain 7 μg, vector DNA 2.3 μg (see Note 7), 10 ligation buffer (Invitrogen) 1 μL, T4 DNA ligase (Invitrogen) 1 μL, and sterile water (total 50 μL) for reaction at 14  C overnight to obtain pCMV-Cx43TC (Fig. 2). 2. Transfect pCMV-Cx43-TC into E. coli: Take out competent cells from a 80  C freezer, dissolve in ice water, add pCMVCx43-TC 20 μL, mix gently for 5 s, maintain for 30 min in ice water, incubate at 42  C for 45 s, and then incubate in ice water for 2 min. Add 900 μL of SOC medium and incubate at 170 rpm, at 37  C for 1 h. Divide the culture to 100 μL and 900 μL fractions and spread on two plates of LB containing Km, respectively, for incubation at 37  C. 3. Analyze the inserted DNA by colony PCR: Prepare a mixture of 5 μL of GoTaq® Green Master Mix, 1 μL of forward primer (50 - GATTTCCAAGTCTCCACCCCA-30 ), 1 μL of reverse primer (50 - TTATGTTTCAGGTTCAGGGGGA-30 ), water 3 μL (total 10 μL). Transfer the DNA of single colony into this mixture by colony pickup with a toothpick. Conduct PCR

pUC ori

PCMV IE Cx43-TC

HSV TK poly A

pCx43-TC 5.2 kb

MCS SV40 poly A

Kan r / Neo r SV40 ori PSV40 e

Fig. 2 pCMV-Cx43-TC vector

f1 P ori

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under the condition: initial denature 94  C 3 min, 30 cycles [denature 94  C 1 min, annealing 56  C 1 min, extension 72  C 2 min], final extension 72  C 2 min. Electrophorese the PCR product on 1% agarose gel to confirm its size (1.5 kb) (see Note 8). 4. Culture of E. coli with correct insertion: Inoculate the colony in LB medium containing 100 μg/mL Km for culture at 37  C at 150 rpm for 12–14 h. Conduct the same procedure as Subheading 3.2, steps 2–4, except that final product is purified pCMV-Cx43-TC. 5. Confirm the sequence of pCMV-Cx43-TC: Prepare a mixture of 0.5 μL of purified plasmid, 0.8 μL of forward primer (50 -G ATTTCCAAGTCTCCACCCCA-30 ), 0.8 μL of reverse primer (50 -TTATGTTTCAGGTTCAGGGGGA-30 ), 4 μL of Pre Mix (Applied Biosystems), and water 3.9 μL (total 10 μL). Conduct PCR under the condition: initial denature 96  C 1 min, 25 cycles [denature 96  C 10 s, annealing 50  C 5 s, extension 60  C 80 s], and final holding at 4  C. Transfer the PCR product to a 1.5 mL microtube, and add ethanol for incubation for 15 min. Centrifuge at 20,000  g at 25  C for 20 min. Wash the precipitate with 70% ethanol, dry, and dissolve with 9 μL Hi-Di formamide. Incubate in the dark at 95  C for 5 min, place on ice for 5 min, and dispense to a microplate for DNA sequence analysis with 3100-Avant Genetic Analyzer (Applied Biosystems). Analyze the homology between cDNA and Cx43 sequences based on BLAST (http://www.ncbi.nlm.nih.gov/ blast/Blast.cgi). 3.5 Transfection of HeLa Cells and FlAsH Staining

1. Dissolve 4 μg of pCMV-Cx43-TC in 250 μL of GMEM. Mix 10 μL of Lipofectamine 2000 and 250 μL of GMEM, and incubate at room temperature for 5 min. Mix these solutions with gentle stirring, and hold at room temperature for 20 min. Add this mixture to a dish of 90% confluent HeLa cells. Refresh the medium after incubation for 4 h. At 20 h later, replace the medium by the selection medium containing 900 μg/μL G418, and culture for 2 successive weeks. 2. Prepare a FlAsh solution (1 μL of FlAsh-EDT2(TRC) and 4 mL of GMEM) and a BAL wash buffer (32 μL of BAL and 8 mL of GMEM). Remove culture medium from a dish of 70–80% confluent HeLa cells after 3–4 days culture, wash twice with PBS, and add 500 nM FlAsH solution for incubation at 37  C for 1 h under 5% CO2. Remove FlAsH solution, wash with 2 mL PBS, and add 2 mL of 100 μM BAL for incubation at 37  C for 10 min under 5% CO2. Repeat the washing twice with PBS and BAL. Wash again with PBS and add 2 mL medium to the dish.

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Fig. 3 Expression of Cx43-TC in HeLa cells. A red circle: a plaque located at the interface of two cells. A yellow circle: a Cx43-TC cluster in the cytosol 3.6 Analysis of GJ Plaque Localization and Gap Junctional Dye Diffusion

1. Prepare a solution of two dyes (9 μL of 8 μg/μL Lucifer yellow and 1 μL of 1 mM Dextran, Texas Red 1 μL). 2. Prepare capillaries for injection: Pull a single channel glass capillary (GF100-78-10, Sutter Instrument Co.) with a lazar puller (P-2000, Sutter Instrument Co.). 3. Observe the expression sites of Cx43-TC in HeLa cells (Fig. 3). 4. Focus a plaque of Cx43-TC localizing at the interface of two cells. Into one of these two cells, inject a solution containing DTR and LY by means of a single-cell manipulation supporting robot (see Note 9). 5. Analyze the dye diffusion to the other cell to confirm the gap junctional permeation of LY with a fluorescent microscope and a camera system (Hamamatsu ORCA-ER and HCImage system) (Hamamatsu Photonics) (Fig. 4).

3.7 Preparation of a Culture Dish with a Coordinate Chip

1. Prepare a right triangle chip depicted in Fig. 5a from a polystyrene plate by cutting with a cutter knife. Take special care to make the right angle part as accurate as possible.

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Fig. 4 Three cases of diffusion properties of transfected HeLa cells. (a1, b1, c1), merge of Cx-TC and DTR; (a2, b2, c2), merge of Cx-TC and LY; (a3, b3, c3), merge of Cx-TC, DTR, and bright field image. Scale bar: 30 μm

2. Disinfect the chip with 70% ethanol, and attach it on the inside of the bottom of a culture dish (3.5 cmϕ) with cyanoacrylate adhesive. 3. Add PBS containing 0.1% gelatin to the dish, place it still for 5 min, and then replace the PBS by 8 mL HeLaM. 4. Define the two sidelines at the right angle as the x and y axes and the right angle corner as the origin (Fig. 5b). 5. Culture HeLa cells in a culture dish (10 cmϕ) with a coordinate chip at 3  105 cells/dish at 37  C under 5% CO2. Select several target cells arbitrarily, and register their x-y coordinates under a fluorescent microscope (Fig. 5b). Inject a solution of 1 mg/mL DTR into the registered target cells of HeLa

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Fig. 5 Schematic diagram of the preparation of an epoxy block containing cell specimens

(Fig. 5c) using a single-cell manipulation supporting robot (see Note 9). Capture fluorescent microscopic images of DTR. 6. Rinse the cells with PBS, and add DMEM containing 2.5% glutaraldehyde and 2% paraformaldehyde to the dish for fixation. Incubate the dish at 20–25  C for 1 h. Wash the dish with distilled water three times, and add 2 mL of 0.1 M PBS containing 1.5% OsO4 to the dish for incubation at 20–25  C for 1 h (Fig. 5d). 7. Wash the dish with distilled water three times, and dehydrate by replacing water by 30%, 50%, 90%, and 100% ethanol successively in this order. Replace ethanol by 10%, 30%, 50%, 70%, and 90% EPON 812 ethanol solutions in this order and finally by 100% EPON 812. Incubate the dish at 20–25  C overnight (Fig. 5e).

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Fig. 6 Engraved shape after the removal of the chip. Observed by light microscopy. O, Ox; y, origin, x axis, and y axis

8. Fill about 80% of the volume of a 1 mL polyethylene capsule (BEEM Capsule, inside diameter 8 mm) with 100% EPON 812. Place the capsule upside down onto the dish, and incubate at 60  C for 48 h to allow the EPON to polymerize (Fig. 5f). 9. Confirm the DTR injected target cells around the x-y coordinates registered initially before incubation at 60  C. 10. Place the dish on a hot plate and heat at 145  C for 5 min (Fig. 5g). Peel off the epoxy block from the dish bottom (Fig. 5h). Place the epoxy block on the hot plate and heated at 145  C for 3 min (Fig. 5i). Separate the right triangle chip with a pair of tweezers from the epoxy block (Fig. 5j). 11. Confirm the origin and x–y axes after the removal of the chip (Figs. 5j and 6). 3.8 Correlation of XY Coordinates Determined by Light and Electron Microscopy

1. Fix the epoxy block on an ultramicrotome and trim it with a razor blade. Slice the epoxy block with a diamond knife to obtain sections with a thickness of 95 nm. Stain the sections with uranyl acetate and then with lead citrate. Observe the sections with an electron microscope (Fig. 7). 2. Analyze the locations of cells registered by light microscopy and observed by electron microscopy (Fig. 8, Table 1). Distance between the cell positions determined by light microscopy and electron microscopy is given by AiBi (i ¼ 1–5) in Table 1. Spatial precision may be given by AiBi/{(OAi + OBi)/ 2}  100%. The average of five cells is 9% (see Note 10).

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Fig. 7 A SEM view of a section prepared from an epoxy block

Fig. 8 Comparative images of No. 1 in Fig. 7. (a) A live image in light microscopy, (b) a live fluorescent image, (c) an image of a fixed cell in epoxy block in light microscopy, (d) an SEM image of a fixed cell in epoxy section. Red arrows in (a)–(d) indicate the same cell

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Table 1 Comparison of cell locations registered by light microscopy and confirmed as the same positions by electron microscopy Ai(x, y) Cell no. (i)

xA

Bi(x, y) yA

xB

yB

xB-XA

yB-yA

OAi (μm)

OBi (μm)

AiBi (μm)

1

760

443

860

495

100

52

880

992

113

2

704

764

766

841

62

77

1039

1138

99

3

550

544

579

565

29

21

774

810

36

4

1168

619

1150

687

18

68

1322

1339

70

5

612

355

664

449

52

94

708

801

107

4

Notes 1. Freezing and thawing of cells: Suspend cells in HeLaM containing 10% DMSO at 2.5  106 cells/0.5 mL, and place it in ice water for 5 min and then 20  C for 30 min. Confirming the freezing of cells, transfer it in a deep freezer at 80  C. After overnight, transfer the cells in a deep freezer at 152  C. Conversely, in thawing, transfer a tube of frozen cells stored at 152  C into a bath at 37  C for rapid thawing. Then suspend it in 5 mL HeLeM. After centrifugation at 350  g at 4  C for 5 min, suspend the precipitate in 4 mL HeLeM, and spread in two dishes. Culture them at 37  C under 5% CO2. 2. Other vectors may be used if they have a promoter. 3. Extract of RNA from mouse ES cells is conducted with ISOGEN (Nippon Gene). Refer to the manufacturer’s protocol. Other cell lines may be used if Cx43 is expressed. 4. Reverse transcription of RNA is conducted with SuperScript II (Invitrogen). Refer to the manufacturer’s protocol. 5. These buffers are used for AgeI and EcoRI, respectively. Consult the manufacturer’s buffer list for other enzymes. 6. Rapid operation is important in order to minimize the injury in DNA by UV illumination. 7. The optimum value of the [vector DNA/insert DNA] ratio depends upon the size of vector DNA. Here, this value is set as 1:3. Variation of this value from 1:2 to 1:10 is recommended. 8. Determine an appropriate density of agarose gel according to the size of the insert DNA. 9. A single-cell manipulation supporting robot enables highthroughput and semiquantitative injection into as small as mouse ES cells. This robot, however, is not commercially

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available. If the size of target cells is as large as 30–40 μm, a commercially available injection machine is applicable. 10. This spatial precision may be a practically acceptable level. If a smaller chip with a more sharp cut right angle is available, much higher precision may be attained.

Acknowledgments We thank Prof. Emer. Hideaki Matsuoka of Tokyo University of Agriculture and Technology for his valuable advice on cell analysis. References 1. Oyamada M, Takebe K, Endo A, Hara S, Oyamada Y (2013) Connexin expression and gap-junctional intercellular communication in ES cells and iPS cells. Front Pharmcol 4:Article 85. https://doi.org/10.3389/fphar.2013. 00085 2. Aasen T (2015) Connexins: junctional and non-junctional modulators of proliferation. Cell Tissue Res 360:685–699. https://doi. org/10.1007/s00441-014-2078-3 3. So¨hl G, Willecke K (2003) An update on connexin genes and their nomenclature in mouse and man. Cell Commun Adhes 10:173–180 4. Axelsen LN, Calloe K, Holstein-Rathlou N-H, Nielsen MN (2013) Managing the complexity of communication: regulation of gap junctions by post-translational modification. Front Pharmacol 4:Article 130 5. Stout RF Jr, Snapp EL, Spray DC (2015) Connexin type and fluorescent protein-fusion tag determine structural stability of gap junction plaques. J Biol Chem 290:23497–23515 6. The´venin AF, Kowal TJ, Fong JT, Kells RM, Fisher CG, Falk MM (2013) Proteins and mechanisms regulating gap-junction assembly, internalization, and degradation. Physiology 28:93–116. https://doi.org/10.1152/ physiol.00038.2012 7. Aasen T, Leithe E, Graham SV, Kameritsch P, Maya´n MD, Mesnil M et al (2019) Connexins in cancer: bridging the gap to the clinic. Oncogene 38:4429–4451. https://doi.org/10. 1038/s41388-019-0741-6 8. Sirnes S, Bruun J, Kolberg M, Kjenseth A, Lind GE, Svindland A et al (2012) Connexin43 acts as a colorectal cancer tumor suppressor and predicts disease outcome. Int J Cancer 131:570–581 9. Wang ZS, Wu LQ, Yi X, Geng C, Li YJ, Yao RY (2013) Connexin-43 can delay early recurrence

and metastasis in patients with hepatitis B-related hepatocellular carcinoma and low serum alpha-fetoprotein after radical hepatectomy. BMC Cancer 13:306. https://doi.org/ 10.1186/1471-2407-13-306 10. Brockmeyer P, Jung K, Perske C, Schliephake H, Hemmerlein B (2014) Membrane connexin 43 acts as an independent prognostic marker in oral squamous cell carcinoma. Int J Oncol 45:273–281 11. Poyet C, Buser L, Roudnicky F, Detmar M, Hermanns T, Mannhard D et al (2015) Connexin 43 expression predicts poor progressionfree survival in patients with non-muscle invasive urothelial bladder cancer. J Clin Pathol 68:819–824 12. Saito M, Asai Y, Imai K, Hiratoko S, Tanaka K (2017) Connexin30.3 is expressed in mouse embryonic stem cells and is responsive to leukemia inhibitory factor. Sci Rep 7:42403. https://doi.org/10.1038/srep42403 13. Hoffmann C, Gaietta G, Zurn A, Adams SR, Terrillon S, Ellisman MH et al (2010) Fluorescent labeling of tetracysteine-tagged proteins in intact cells. Nat Prot 5:1666–1677 14. Saito M, Imai K, Koyama M (2016) A tetracysteine-tag and HeLa cell system for the dynamic analysis of the localization and gating properties of a specific connexin isoform. Electrochemistry 84(5):299–301. https://doi. org/10.5796/electrochemistry.84.299 15. Haraguchi T, Osakada H, Koujin T (2015) Live CLEM imaging to analyze nuclear structures at high resolution. In: Nakagawa S, Hirose T (eds) Nuclear bodies and noncoding RNAs—methods and protocols, methods in molecular biology 1262, Springer Protocols. Humana Press, Totowa, pp 89–103 16. Peddie CJ, Domart MC, Snetkov X, O’Toole P, Larijani B, Way M et al (2017)

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Correlative super-resolution fluorescence and electron microscopy using conventional fluorescent proteins in vacuo. J Struct Biol. pii: S1047-8477(17)30093-X. https://doi.org/ 10.1016/j.jsb.2017.05.013 17. Ariotti N, Hall TE, Parton RG (2017) Correlative light and electron microscopic detection of GFP-labeled proteins using modular APEX. Methods Cell Biol 140:105–121. https://doi. org/10.1016/bs.mcb.2017.03.002 18. Saito M, Hiratoko S, Fukuba I, Tate S, Matsuoka H (2018) Use of a right triangle chip and its engraved shape as a transferrable x-y coordinate system from light microscopy to electron microscopy. Electrochemistry 86(1):6–9

19. Yamada Y, Yamaguchi N, Ozaki M, Shinozaki Y, Saito M, Matsuoka H (2008) Instant cell recognition system using microfabricated coordinate standard chip useful for combinable cell observation with multiple microscopic apparatus. Microsc Microanal 14:236–242. https://doi.org/10.1017/ S1431927608080252 20. Saito M, Matsuoka H (2010) Semiquantitative analysis of transient single-cell gene expression in embryonic stem cells by femtoinjection. In: Zhang B (ed) RNAi and microRNA-mediated gene regulation in stem cells, Methods in molecular biology 650. Humana Press, Totowa, pp 155–170. https:// doi.org/10.1007/978-1-60761-769-3_13

Methods in Molecular Biology (2021) 2346: 207–214 DOI 10.1007/7651_2020_291 © Springer Science+Business Media New York 2020 Published online: 17 June 2020

Chemical and Voltage Gating of Gap Junction Channels Expressed in Xenopus Oocytes Camillo Peracchia Abstract In most tissues, cells in contact with each other directly intercommunicate via cell-to-cell channels aggregated at gap junctions. Direct cell-to-cell communication provides a fundamental mechanism for coordinating many cellular functions in mature and developing organs, as it enables free exchange of small cytosolic molecules. Gap junction channels are regulated by a chemical gating mechanism sensitive to cytosolic calcium concentration [Ca2+]i in the nanomolar range mediated by Ca2+-activated calmodulin (CaM). Evidence for the relevance of chemical regulation of gap junctional communication to cell function in health and disease prompted the development of methodologies aimed at quantitatively monitoring channel gating. A widely used method is the double voltage clamp of Xenopus laevis oocytes. Basically, this method involves pairing at the vegetal pole devitellinized oocytes in a conical well of a culture dish, inserting in each of them a current and a voltage microelectrode, establishing double voltage clamp and measuring junctional conductance (Gj) from voltage and current records. Key words Gap junctions, Connexins, Channel gating, Cell-to-cell communication, Calcium, pH, Calmodulin, Double voltage clamp

1

Introduction In normal functional circumstances, most gap junction channels are in an open state, enabling neighboring cells to freely exchange charged and neutral cytosolic molecules as heavy as about 1 kDa [1–3]. These channels, however, like other membrane channels, possess gates that when activated reduce channel permeability. Both chemical and trans-junctional voltage (Vj)-sensitive gating mechanisms are present. While activated Vj gates close the channels by ~80%, chemical gates close them completely [4]. Activation of chemical gates causes neighboring cells to uncouple from each other electrically and metabolically. In early studies, uncoupling was believed to be just a protective mechanism that enables cells to survive the damage or death of neighboring cells. In contrast, recent evidence for channel permeability regulation by cytosolic [Ca2+]i in the nanomolar range [5–9] suggests the role of calmodulin in chemical gating [5, 7, 10–12] and indicates that the fine modulation of channel gating plays a role in normal cellular

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functions as well. To understand how gap junction communication is physiologically controlled, starting in the late 1990s, we have measured junctional conductance (Gj) of various connexins and connexin-mutant channels in Xenopus oocyte pair by double voltage clamp [13]. With this method we have tested the channels’ gating behavior by various approaches including inhibition of calmodulin (CaM) expression, expression of CaM mutants, identification of connexin domains relevant for gating, expression of connexin mutants, testing of Vj-gating sensitivity to CO2-induced acidification, and testing repeated application of large Vj gradients, among others, rev. in [5, 6]. Oocytes provide an excellent model system for many reasons:

2

They allow one to perform long experiments with the double voltage clamp technique.

l

Are continuously produced.

l

Oocytes in different stages are always present in ovaries.

l

The antibiotic nature of the Xenopus’ natural skin secretion prevents postoperative infection.

l

Since a Xenopus can donate oocytes at least four times, 20–30 frogs are enough for performing at least two experiments per week.

Materials

2.1 Solutions Used During Oocyte Preparation and in Electrophysiological Experiments

3

l

The Ca-free OR2 solution with 2 mg/ml collagenase type 1A (Sigma Chemical Co., St Louis, MO, USA) contains (in mM): NaCl 82.5, KCl 2, MgCl2 1, and 1,4-(2-hydroxyethyl)-1-piperazineethanesulfonic acid (HEPES) 5 (pH 7.6 with NaOH). The standard ND96 solution contains (in mM): NaCl 96, KCl 2, CaCl2 1.8, MgCl2 1, and HEPES 5 (pH 7.6 with NaOH). The Cl-free saline contains (in mM) NaOH 75, KOH 10, Ca (OH)2 4, Mg(OH)2 5, and 3-(N-morpholino)propanesulfonic acid (MOPS) 10, adjusted to pH 7.2 with methanesulfonic acid.

Methods

3.1 Oocyte Preparation

Oocytes are surgically removed from the abdomen of adult Xenopus laevis females. Following anesthesia by immersion in dechlorinated water containing 3.7 g of tricaine/l, a 1-cm incision is made through the skin and muscle layers of the lateral abdominal wall. Approximately one eighth of the ovarian lobe is pulled through the incision and cut off and placed in culture medium. After the ovarian lobe retracts back into the abdomen, muscle wall and skin are closed with four to five stitches (4-O chromic gut) of the

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Fig. 1 Mattress-type, vertical Donati’s stitching (a). This technique allows precise eversion of the cut’s edges (b)

“mattress-type” stitching, also known as “vertical Donati’s stitching”1 (Fig. 1a, b) (see Note 1). The Xenopus recovers from anesthesia in a basin containing shallow water (see Note 2). Stage V or VI oocytes are defolliculated in 2 mg/ml collagenase type 1A (Sigma Chemical Co.) in Ca-free OR2 solution and allowed to recover overnight in standard ND96 solution at 18  C. If needed, the oocyte’s expression of the endogenous connexin (Cx38) is blocked by injecting antisense oligonucleotides (see in the following), so that other connexin genes can be expressed. 3.2 Inhibition of Native-Connexin Expression

1

Xenopus oocytes express a native connexin, Cx38, which can form not only homotypic channels but also heterotypic (and possibly heteromeric) channels with several mammalian connexins. Therefore, for testing the function of exogenous connexins and their mutants without interference from Cx38, the expression of this connexin needs to be inhibited. This can be accomplished by injecting defolliculated oocytes with 46 nl of 0.25 mg/ml antisense oligonucleotide complementary to endogenous Xenopus Cx38. The antisense oligonucleotide blocks completely the endogenous junctional communication within 48 h. Cx38 antisense oligonucleotides complementary to different Cx38 domains are used: either 50 -GCTTTAGTAATTCCCATCCTGCCATGTTTC-30 (commencing at nucleotide (nt) 5 of Cx38’s cDNA sequence [14] or 50 -AGCAGAAGAGTATACTTCTGTTTGT-3 (commencing at nt 363 of Cx38’s cDNA sequence).

The Donati’s stitching is named after its inventor—the renowned Italian surgeon Mario Donati (1879–1946).

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3.3 Inhibition of Calmodulin (CaM) Expression

For experiments testing the effects of inhibiting CaM expression, oocytes injected 24-h earlier with oligonucleotides antisense to Cx38 are injected with oligonucleotides antisense to the two CaM mRNAs expressed by Xenopus oocytes (46 nl, 1.12 mg/ ml) [13].

3.4 Expression of CaM Mutant

Oocytes previously injected with antisense oligonucleotide complementary to endogenous Xenopus Cx38 [10, 15] are injected at the vegetal pole with cRNA of the CaM mutant CaMCC and incubated overnight at 18  C [10]. In CaMCC, the N-terminal EF hand pair (residues 9–75) is replaced by a duplication of the C-terminal pair (residues 82–148). The oocytes are then reinjected 24 h later with Cx32 cRNA and homotypically paired 724 h later.

3.5 Oocyte Injection of Connexin cRNAs and Oocyte Pairing

Forty-eight to seventy-two hours after the injection of antisense oligonucleotides complementary to endogenous Xenopus Cx38, 46 nl of either a wild type connexin or a connexin mutant (0.04–0.2 mg/ml) cRNA is injected into oocytes at the vegetal pole and the oocytes are incubated overnight at 18  C. The oocytes are mechanically stripped of their vitelline layer in a hypertonic medium by pinching the vitelline layer with two fine forceps (see Note 3) and paired at the vegetal poles in a conical well of a Petri dish containing conical wells (Fig. 2A) (Falcon polystyrene microplates 96-well) filled with ND96. The Petri dish contains an epoxyglued square segment (Fig. 2A, b) of the Falcon microplate (Fig. 2A, a). The oocyte pairs are studied electrophysiologically 0.5–3 h after pairing.

3.6 Measurement of Gap Junctional Conductance by Double Voltage Clamp

Microelectrodes are pulled from borosilicate glass capillaries 1.2 mm (o.d.) 0.68 mm (i.d.), by means of a micropipette puller. The microelectrodes, filled with a 3 M KCl solution, have a resistance of 0.5–1 MΩ in ND96. The bath is grounded with a silversilver chloride reference electrode connected to the perfusion chamber via an agar bridge. The experiments are performed using the standard double voltage-clamp procedure for measuring junctional conductance (Gj) [16, 17]. Following the insertion of a current and a voltage microelectrode in each oocyte (Fig. 2B), both oocytes are initially voltage clamped to the same holding potential, Vm1 ¼ Vm2 (usually 20 mV), so that no junctional current flows at rest (Ij ¼ 0). A Vj gradient is created by imposing a +20 mV voltage pulse (V1) of 2 s duration every 10 or 30 s to oocyte 1 while maintaining V2 at Vm; thus, Vj ¼ V1. The negative-feedback current (I2), injected by the clamp amplifier in oocyte 2 for maintaining V2 constant at Vm, is used for calculating Gj, as it is identical in magnitude to the junctional current (Ij), but of opposite sign (Ij ¼ I2); Gj ¼ Ij/Vj (Ohm’s law). Pulse generation and data acquisition are performed by means of a computer equipped with pCLAMP software and A/ D-D/A interface (Molecular Devices, LLC. San Jose, CA).

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Fig. 2 Stripped oocytes are paired at the vegetal poles in a conical well of a Falcon microplate fragment epoxy-glued in a Petri dish (A, a and b). Four microelectrode are inserted in the oocytes to perform double voltage clamp (B). A Vj gradient is created by imposing a +20 mV voltage pulse (V1) to oocyte 1, while maintaining V2 at Vm (Vj ¼ V1). The negative feedback current (I2), injected in oocyte 2 for maintaining V2 at Vm, is used for calculating Gj (Ij ¼ I2). Gj ¼ Ij/Vj (Ohm’s law) 3.7 Measurement of Trans Junctional Voltage Gating

For studying voltage dependence of junctional conductance, each oocyte of the pair is first voltage clamped at 20 mV. Voltage pulses of 20 mV (increased step wise up to 120 mV maximum) and 20 s duration are then applied every 45 s to either oocyte of the pair while maintaining the other at 20-mV. To illustrate the relationship between steady-state Gj (Gj ss) and Vj, the normalized Gj (Gj ss/Gj max) is plotted with respect to Vj. The curve is fit to a two-state Boltzmann distribution of the form (Gj ss  Gj min)/ (Gj max  Gj ss) ¼ exp.[A(Vj  V0)], where V0 is the Vj value at

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which the voltage-sensitive conductance is one half the maximal value and Gj min is the theoretical minimum normalized Gj. A ¼ nq/kT is a constant that expresses voltage sensitivity in terms of number of equivalent gating charges, n, moving through the entire applied field, where q is the electron charge, k is the Boltzmann constant, and T is the temperature in degrees Kelvin. 3.8 Oocyte Chamber Perfusion and Uncoupling Protocol

The oocyte chamber is continuously perfused at a flow rate of 0.6ml/min by a peristaltic pump. The perfusion solution is ejected by a 22-gauge needle placed near the edge of the conical well containing the oocyte pair. The level of the solution in the chamber is maintained constant by continuous suction. Electrical uncoupling of oocyte pairs is induced by a 3- to 15-min perfusion (0.6-ml/min) of saline continuously gassed with 100% CO2. A Cl free saline (Cl replaced with methane sulfonate) is used because the opening of Ca2+-activated Cl-channels during exposure to 100% CO2 causes an increase in membrane current that may interfere with measurements of junctional current [13]. For some reasons, probably related to seasonal variations in Xenopus physiology, this phenomenon is more pronounced in the summer and early fall seasons.

3.9 Bubble-Trap Device

The removal of gas bubbles from perfusion solutions gassed with 100% CO2 or other gasses is accomplished by driving the perfusion solution through a glass connector shaped as an inverted “Y” (Fig. 3) (see Note 4). The solution enters the connector from one of the two branches of the Y-connector that points downward and exits through the other branch with similar downward orientation.

Fig. 3 Device designed to remove gas bubbles from perfusion solutions gassed with 100% CO2 or other gasses. As the solution reaches the center of the connector (the intersection among the three branches) it is exposed to air contained in the syringe. This enables gas bubbles carried by the inflowing solution to escape into the air-filled syringe cavity

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The branch pointing upward is connected to an empty syringe whose piston is partly withdrawn to create an air volume of 40–50 ml. The solution reaching the center of the connector (the intersection among the three branches) is exposed to air contained in the syringe. This enables gas bubbles carried by the inflowing solution to escape into the air-filled syringe cavity. The level of the solution exposed to air can be regulated by withdrawing or depressing the syringe piston.

4

Notes 1. Since the Xenopus skin is very thin, with conventional stitching it is difficult to prevent infolding of the cut’s edges. Therefore, it is better to use the mattress-type, vertical Donati’s stitching (Fig. 1a), because this stitching technique allows precise eversion of the cut’s edges (Fig. 1b). 2. It is important that the water is dechlorinated because chlorine would damage the Xenopus’ skin. Also, one should make sure that the Xenopus’ nose always remains above water until she can move well inside the basin on her own. For fully evaporate chlorine from 10 gallons of tap water at room temperature it takes 4–5 days. One way to remove chlorine faster is to boil the tap water, as it will take 68 min to completely remove it. 3. The forceps need to be carefully filed to create rounded, smooth, tips in order to prevent oocyte damage during the vitelline-membrane stripping procedure. 4. Perfusion solutions continuously gassed with CO2 or other gases may generate bubbles that when ejected near the oocyte pair cause vibrations and may result in irreversible oocyte damage. To prevent it we have constructed a “bubble-trap” device (Fig. 3) designed to remove gas bubbles before the perfusion solutions reach the oocyte bath. The “bubble-trap” device works well only with connectors made of glass or other materials that, like glass, provide hydrophilic surfaces. The internal surface of the glass connector needs to be kept clean to remain hydrophilic. This is accomplished by cleaning the perfusion system with methanol at the end of each experiment.

References 1. Loewenstein WR (1975) Permeable junctions. Cold Spring Harb Symp Quant Biol 40:49–63 2. Evans WH (2015) Cell communication across gap junctions: a historical perspective and current developments. Biochem Soc Trans 43 (3):450–459

3. Evans WH, Martin PE (2002) Gap junctions: structure and function (review). Mol Membr Biol 19(2):121–136 4. Bukauskas FF, Peracchia C (1997) Two distinct gating mechanisms in gap junction channels: CO2-sensitive and voltage-sensitive. Biophys J 72(5):2137–2142

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5. Peracchia C (2020) Calmodulin-mediated regulation of gap junction channels. Int J Mol Sci 21(2):485 6. Peracchia C (2019) Gap junction structure and chemical regulation. Direct calmodulin role in cell-to-cell channel gating. Academic Press, London 7. Peracchia C (2004) Chemical gating of gap junction channels; roles of calcium, pH and calmodulin. Biochim Biophys Acta 1662 (12):61–80 8. Lazrak A, Peracchia C (1993) Gap junction gating sensitivity to physiological internal calcium regardless of pH in Novikoff hepatoma cells. Biophys J 65(5):2002–2012 9. Xu Q, Kopp RF, Chen Y, Yang JJ, Roe MW, Veenstra RD (2012) Gating of connexin 43 gap junctions by a cytoplasmic loop calmodulin binding domain. Am J Physiol Cell Physiol 302(10):C1548–C1556 10. Peracchia C, Sotkis A, Wang XG, Peracchia LL, Persechini A (2000) Calmodulin directly gates gap junction channels. J Biol Chem 275 (34):26220–26224 11. Peracchia C, Bernardini G, Peracchia LL (1983) Is calmodulin involved in the

regulation of gap junction permeability? Pflugers Arch 399(2):152–154 12. Zou J, Salarian M, Chen Y, Veenstra R, Louis CF, Yang JJ (2014) Gap junction regulation by calmodulin. FEBS Lett 588(8):1430–1438 13. Peracchia C, Wang X, Li L, Peracchia LL (1996) Inhibition of calmodulin expression prevents low-pH-induced gap junction uncoupling in Xenopus oocytes. Pflugers Arch 431 (3):379–387 14. Rubin JB, Verselis VK, Bennett MVL (1992) Molecular analysis of voltage dependence of heterotypic gap junctions formed by connexins 26 and 32. Biophys J 62:183–195 15. Sotkis A, Wang XG, Yasumura T, Peracchia LL, Persechini A, Rash JE, Peracchia C (2001) Calmodulin colocalizes with connexins and plays a direct role in gap junction channel gating. Cell Commun Adhes 8(4–6):277–281 16. Spray DC, Harris AL, Bennett MV (1981) Equilibrium properties of a voltage-dependent junctional conductance. J Gen Physiol 77 (1):77–93 17. Neyton J, Trautmann A (1985) Single-channel currents of an intercellular junction. Nature 317(6035):331–335

Methods in Molecular Biology (2021) 2346: 215–223 DOI 10.1007/7651_2020_322 © Springer Science+Business Media New York 2020 Published online: 20 September 2020

The Analysis of Gap Junctional Intercellular Communication Among Osteocytes in Chick Calvariae by Fluorescence Recovery After Photobleaching Ziyi Wang, Yoshihito Ishihara, and Hiroshi Kamioka Abstract This chapter describes the use of fluorescence recovery after photobleaching (FRAP) for analyzing gap junctional intercellular communication (GJIC) among osteocytes in chick calvariae by confocal laser scanning microscope. Keywords Osteocytes, Gap-junctional intercellular communication, Fluorescence recovery after photobleaching, One-phase exponential association equation

1

Introduction Cell-cell communication is crucial for the cell as a member of a tissue or an organ. Osteocytes are the most abundant cells in bone, forming a three-dimensional (3D) structural network by connecting with each other through long, slender cell processes. They help contribute to synthesize, resorb, and repair skeletal tissue by directing the differentiation and activity of surface osteoblasts and osteoclasts. Blockade of these networks among bone cells diminishes the bone quality and leads to skeletal fragility [1].

1.1 Osteocyte Network

Gap junctional intercellular communications (GJICs) play an important role in which bone cells coordinate their actions. Small ions, molecules, and second messengers can be directly exchanged between bone cells via the aqueous channels formed by gap junctions. Therefore, GJICs form an efficient interconnected cellular network that results in the functional and efficient coordination among bone cells. The gap junctions presented in bone cells are able to sense forces, since the mechanical simulation of osteocytes increases the expression of connexins in vitro and in vivo, presumably creating enhanced connections with neighboring cells to strengthen the transmission of mechanical information within the osteocyte network [2]. Our previous study showed that extracellular pH, extracellular calcium, and parathyroid hormone

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significantly altered the rate of fluorescence recovery after photobleaching [3]. Indeed, the maturation of osteocytes affects the rate of recovery of fluorescence, and the speed of dye diffusion may differ as well [4]. 1.2 Fluorescence Recovery After Photobleaching

2

Compared with the use of fluorescence dye via microinjection or scrape loading, fluorescence recovery after photobleaching (FRAP) is noninvasive and easier and faster to perform. In the FRAP technique, we use the chemical and biophysical properties (such as the size) of tracers without a metabolic role to define and quantify the communication capacity. In general, it was assumed that the choice of dye would not significantly influence the fluorescence redistribution. A previous study showed that the rate of fluorescence recovery did not differ significantly among different dyes, even when their fluorescence kinetic profiles were obviously different [5].

Materials Animals

Lohmann LSL-classic embryonic chicks were obtained from Lohmann Tierzucht (Cuxhaven, Germany).

2.2 General Reagents

1. Culture medium: alpha minimal essential medium (α-MEM) without antibiotics.

2.1

2. Fetal bovine serum (FBS). 3. Calcein acetoxymethyl ester (calcein AM; Molecular Probes Inc., Eugene, OR, USA), a membrane-permeable dye. Once calcein AM permeates the cytoplasm, it is hydrolyzed by esterase to calcein (molecular weight ¼ 622), which remains inside the cell. 2.3

Equipment

1. Egg incubator. 2. Compact stereo microscopes for bone fragment preparation (Olympus SZ61 Stereo Microscope 0.67x-4.5x; Olympus, Tokyo, Japan). 3. We routinely use a FLUOVIEW FV500 confocal laser scanning microscopy (CLS) system (Olympus) equipped for differential interference contrast (DIC) microscopy. For the FRAP experiment with calcein staining, 488 nm light was emitted by an Argon laser (50 mW on max). The CLS microscopy system was coupled to an upright microscope (IX-70; Olympus) with a x60 or x40 (NA ¼ 1.4 or 1.35) oil immersion objective lens. The scanning rate was 2.71 s/scan for an 8-bit image, 512  512 pixels in size.

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4. Glass-bottom (0.16–0.19 mm thick) plastic dish (Matsunami, Osaka, Japan). 5. Disposable biopsy punch with a plunger system (Kai Industries Co., Gifu, Japan).

3

Methods

3.1 Tissue Collection and Calcein AM Loading

1. Calvariae were obtained from 16-day-old embryonic chicks and washed with α-MEM to remove nonadherent cells. 2. The calvariae were trimmed into 3  3 mm pieces. The average thickness of the sample ranged from 60 to 80 μm (see Note 1). 3. The bone fragments were loaded with 5.0 μM calcein AM in α-MEM for 15 min at 37 C and in 5% CO2. 4. The bone fragments were washed using α-MEM three times to remove excess dye. 5. The bone fragments were incubated for 45 min in α-MEM with 2% FBS at 37 C and in 5% CO2. 6. The bone fragments were washed using α-MEM three times and then placed in glass-bottom plastic dish with α-MEM only (no FBS see Note 2). 7. The bone fragments were held in place by a coverslip secured using adhesive grease.

3.2 FRAP Experiment on an Osteocyte

1. An osteocyte was located using the x60 objective (see Note 3). 2. To capture images, the nominal speed “Fast” (1.66 s/scan) was used, which takes 1.7 s to finish a scan. Different pinhole sizes (50–100 μm) were tried in order to obtain strong fluorescence. When taking images, we used a low laser transmission, e.g., 1–5%, to avoid photobleaching the entire image. 3. The target osteocyte was selected using the region-of-interest (ROI) tool. For the FRAP experiment, we used control images before bleaching and then bleached the ROI 1 time (1 scan) at nominal 50% laser transmission (acousto-optic tunable filter [AOTF] ¼ 50% in this experiment) and captured a series of images immediately after bleaching. For this experiment, images were captured every 30 s for 300 s after bleaching (see Note 4). 4. The images were then saved.

3.3

Data Analyses

1. Images were opened with the ImageJ software program (see Note 5). 2. The stack of images was aligned using the align tool (“plugins” ! “Template Matching” ! “align slices in stack”; https://sites.google.com/site/qingzongtseng/template-

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matching-ij-plugin) in ImageJ, so that the target osteocytes did not float, instead remaining in the same position on the image (see Note 6). 3. For double normalization, the average intensity at each imaging time point was measured for three ROIs: the bleached target osteocyte (It), all other osteocytes in the image field (Tt) as controls, and a random non-fluorescent region outside of all of the cells for background subtraction (BG) using the “Time Series Analyzer” tool of ImageJ (“plugins” ! “Time Series Analyzer”; https://imagej.nih.gov/ij/plugins/timeseries.html). The fluorescence intensity of the target osteocyte (F) was normalized as follows [6]:  T prebleach  BG ðI t  BGÞ  Ft ¼ ðT t  BGÞ I prebleach  BG

ð1Þ

4. The recovery of fluorescence within a bleached osteocyte (Rt) was calculated using the following equation [3]: Rt ¼ ½ðF t  F 0 Þ=ðF i  F 0 Þ  100 ð%Þ

ð2Þ

The percent recovery was defined as the fraction of molecules that were replaced during the time course of the experiment. Ft is the normalized fluorescence intensity at the time (t) after photobleaching by Eq. 1. F0 is the normalized fluorescence immediately after photobleaching. Fi is the initial fluorescence intensity before photobleaching (see Note 7). 5. Recovery curves were analyzed to determine the passive transport of fluorescent dyes through osteocyte dendritic processes connected by gap junctions. The kinetics followed the following equation [7]: R p  Rt ¼ e kt Rp  R0

ð3Þ

where Rp, R0, and Rt are the recovery of fluorescence in the bleached target osteocyte at the plateau (equilibrium), zero time, and time (t) after bleaching, respectively. Equation 3 can be rearranged as Rt ¼ R0 + (Rp  R0)  (1  ekt), also known as a one-phase exponential association equation. The parameters k and Rp were estimated using the curve fit function in the Graphpad Prism software program or other similar software programs (Fig. 1).

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Fig. 1 FRAP measurements of calcein fluorescence in an osteocyte embedded in isolated chick calvariae. (a) Time-lapse confocal images of a photobleached osteocyte. (b) FRAP curves of calcein fluorescence over a 450-s period. The average fluorescence before photobleaching was counted as 100%. In this example, the mobile fraction (MF) was 44.98%, which is estimated by the predicted plateau from the one-phase exponential association equation; the immobile fraction (IF) is 55.02%, which is estimated by IF ¼ 1 – MF; the constant k is 0.00788, and the half-time is estimated by the equation ln(2)/k to be 87.97 s. This figure was modified from our published work [4] with permission from Elsevier

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6. By ignoring the recovery of fluorescence during the photobleaching period, we were able to consider the Rp as the mobile fraction ( fm), and the immobile fraction ( fi) could then be calculated using the following equation: fi ¼ 1  fm. The rate constant k is positively related to the permeability coefficient of the fluorescence dye (calcein AM in this experiment); however, more morphological information is required to estimate this permeability coefficient, since the cell bodies of osteocytes are not connected directly by gap junctions (see Note 8). The time for 50% plateau recover (half-time) was estimated using the equation ln(2)/k.

4

Notes 1. The calvarial bone on the top of the head (parietal part) is very thin, easy to fold, and is almost entirely composed of young osteocytes (Fig. 2b–d). The bone had to be carefully flattened under a microscope to ensure that only a single layer was captured. In addition to using the parietal part of the calvarial bone, we also used the skull base frontal bone, which is located under and in front of the chick brain and behind the eye (Fig. 2e–h). This part of the skull base frontal bone is much harder and thicker than the calvarial portions. We previously reported the growth rate in this bone section [4]. In this way we were able to locate osteocytes with different “ages” based on our logarithmic growth model, since the size of this hard and thick bone can be easily measured. 2. FBS 2–5% was allowed to be added to the α-MEM in Subheadings 3.1.3–3.1.6. Based on the authors’ personal observations, the presence of FBS may result in better calcein AM loading for the skull base bone segments. However, FBS may cause variation in the FRAP results, especially when treating bone fragments with PTH, gap junction inhibitor, or different extracellular pH or calcium concentrations. Therefore, performing a serum test to determine the best concentration of FBS for chick samples is recommended. 3. When using the skull base frontal bone and searching a mature osteocyte, a x40 lens is recommended, since this part of the bone is much thicker than other parts and the mature osteocytes are located at a deeper point. 4. The AOTF is 100% for the skull base part, as it is much denser and thicker than the parietal part. The best laser power should be determined for different CLS systems. At the very least, the laser power should be set so that the target cell is invisible to the naked eye. Care should be practiced here, as a high laser power can damage the target cell. The time interval and period of the

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Fig. 2 Different sites for the isolation of bone fragments on chick calvariae. (a) The blue area indicates the parietal part, and the red area indicates the skull base part (frontal bone). (b) Appearance of the parietal part of chick calvariae. The isolated calvarium grows in the direction of the arrow (from the orbital ridge to the parietal site). Texas Red-X conjugated phalloidin-labeled (c) young and (d) mature osteocytes from the parietal part of the chick calvariae. (e) Appearance of the skull base part of the chick calvariae. (f) The preparation of bone fragments. Calcein AM-stained (g) young and (h) mature osteocytes from the skull base part of the chick calvariae. Bars in (c) and (d) ¼ 30 μm, in (e) and (f) ¼ 1 mm, and in (g) and (h) ¼ 20 μm. This figure was modified from our published work [4, 9] with permission from Elsevier and Cambridge University Press

time-lapse images may vary. In our experience, the maximal recovery rate is reached after 5 min on average, and the time interval should be kept at