Exocytosis and Endocytosis: Methods and Protocols 1071610430, 9781071610435

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Methods in Molecular Biology 2233

Florence Niedergang Nicolas Vitale Stéphane Gasman Editors

Exocytosis and Endocytosis Methods and Protocols

METHODS

IN

MOLECULAR BIOLOGY

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

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

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

Exocytosis and Endocytosis Methods and Protocols

Edited by

Florence Niedergang Department of Infection, Immunity & Inflammation, Institut Cochin (Institut National de la santé et de la recherche médicale, Centre National de la Recherche Scientifique and Université de Paris), Paris, France

Nicolas Vitale Centre National de la Recherche Scientifique, Institut des Neurosciences Cellulaires et Intégratives, Université de Strasbourg, Strasbourg, France

Stéphane Gasman Centre National de la Recherche Scientifique, Institut des Neurosciences Cellulaires et Intégratives, Université de Strasbourg, Strasbourg, France

Editors Florence Niedergang Department of Infection, Immunity & Inflammation Institut Cochin (Institut National de la sante´ et de la recherche me´dicale, Centre National de la Recherche Scientifique and Universite´ de Paris) Paris, France

Nicolas Vitale Centre National de la Recherche Scientifique Institut des Neurosciences Cellulaires et Inte´gratives Universite´ de Strasbourg Strasbourg, France

Ste´phane Gasman Centre National de la Recherche Scientifique Institut des Neurosciences Cellulaires et Inte´gratives Universite´ de Strasbourg Strasbourg, France

ISSN 1064-3745 ISSN 1940-6029 (electronic) Methods in Molecular Biology ISBN 978-1-0716-1043-5 ISBN 978-1-0716-1044-2 (eBook) https://doi.org/10.1007/978-1-0716-1044-2 © Springer Science+Business Media, LLC, part of Springer Nature 2021 This work is subject to copyright. All rights are reserved by the Publisher, whether the whole or part of the material is concerned, specifically the rights of translation, reprinting, reuse of illustrations, recitation, broadcasting, reproduction on microfilms or in any other physical way, and transmission or information storage and retrieval, electronic adaptation, computer software, or by similar or dissimilar methodology now known or hereafter developed. The use of general descriptive names, registered names, trademarks, service marks, etc. in this publication does not imply, even in the absence of a specific statement, that such names are exempt from the relevant protective laws and regulations and therefore free for general use. The publisher, the authors, and the editors are safe to assume that the advice and information in this book are believed to be true and accurate at the date of publication. Neither the publisher nor the authors or the editors give a warranty, expressed or implied, with respect to the material contained herein or for any errors or omissions that may have been made. The publisher remains neutral with regard to jurisdictional claims in published maps and institutional affiliations. This Humana imprint is published by the registered company Springer Science+Business Media, LLC, part of Springer Nature. The registered company address is: 1 New York Plaza, New York, NY 10004, U.S.A.

Preface Eukaryotic cells have evolved from primitive cells by expanding the original plasma membrane from a protective barrier that preserves the unique chemical composition and architecture of the cell interior to an exquisite hub serving as a front line for communication with its environment. These communications are conveyed by messages partly mediated by vesicular membrane events entering and exiting this hub. For instance like planes in airports, vesicles are constantly budding off the plasma membrane to enter the cytoplasm or arriving from the cytoplasm before fusing with the plasma membrane. These processes are called endocytosis and exocytosis, respectively. From the cellular to the organism level, endocytosis and exocytosis mediate crucial physiological functions in normal and pathological conditions. In consequence, a better understanding of the fundamental mechanisms that govern these processes is of great importance for both basic and applied research. The chapters in this book cover methods to characterize the different pathways of endocytosis, real-time imaging of endocytic steps, endocytosis in model organisms, superresolution methods to follow proteins involved in exocytosis, and specific protocols for exocytosis in specialized cells such as neutrophils, adipocytes, or neuroendocrine cells. Written in the highly successful Methods in Molecular Biology series format, the chapters include introductions to their respective topics, lists of the necessary materials and reagents, step-by-step, readily reproducible laboratory protocols, and tips on troubleshooting and avoiding known pitfalls. Cutting edge and clearly written, Exocytosis and Endocytosis is a valuable resource for researchers in the fields of not only cell biology but also neurology, immunology, or oncology, who are interested in studying protein trafficking and signal regulation. Paris, France Strasbourg, France Strasbourg, France

Florence Niedergang Ste´phane Gasman Nicolas Vitale

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

PART I

v ix

ENDOCYTOSIS

1 Stoichiometry of Receptors at the Plasma Membrane During Their Endocytosis Using Total Internal Reflection Fluorescent (TIRF) Microscopy Live Imaging and Single-Molecule Tracking. . . . . . . . . . . . . . 3 Laura Salavessa and Nathalie Sauvonnet 2 Measuring Endocytosis During Proliferative Cell Quiescence. . . . . . . . . . . . . . . . . 19 Claudia Hinze, Kieran McGourty, and Emmanuel Boucrot 3 Measurements of Compensatory Endocytosis by Antibody Internalization and Quantification of Endocytic Vesicle Distribution in Adrenal Chromaffin Cells . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 43 Mara Ceridono, Sylvette Chasserot-Golaz, Nicolas Vitale Ste´phane Gasman, and Ste´phane Ory 4 Quantitative Methods to Study Endocytosis and Retrograde Transport of Cargo Proteins. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 53 Massiullah Shafaq-Zadah, Estelle Dransart, and Ludger Johannes 5 High-Content Drug Discovery Screening of Endocytosis Pathways . . . . . . . . . . . 71 David A. Cardoso, Ngoc Chau, and Phillip J. Robinson 6 Methods for Monitoring Endocytosis in Astrocytes . . . . . . . . . . . . . . . . . . . . . . . . . 93 Maja Potokar, Jernej Jorgacˇevski, and Robert Zorec 7 Monitoring Activity-Dependent Bulk Endocytosis in Primary Neuronal Culture Using Large Fluorescent Dextrans. . . . . . . . . . . . . . . . . . . . . . . . 101 Michael A. Cousin and Karen J. Smillie

PART II

EXOCYTOSIS

8 Quantitative Flow Cytometry-Based Assays for Measuring Constitutive Secretion . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 115 David E. Gordon, Amber S. Shun-Shion, Asral W. Asnawi, and Andrew A. Peden 9 High-Throughput Screening for Insulin Secretion Modulators . . . . . . . . . . . . . . . 131 Michael A. Kalwat 10 Different Approaches to Record Human Sperm Exocytosis . . . . . . . . . . . . . . . . . . 139 Laila Suhaiman, Karina Noel Altamirano, Alfonsina Morales, and Silvia Alejandra Belmonte

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Bovine Chromaffin Cells: Culture and Fluorescence Assay for Secretion . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Tamou Thahouly, Emeline Tanguy, Juliette Raherindratsara Marie-France Bader, Sylvette Chasserot-Golaz, Ste´phane Gasman, and Nicolas Vitale Measurement of Exocytosis in Genetically Manipulated Mast Cells . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Ofir Klein, Nurit P. Azouz, and Ronit Sagi-Eisenberg Super-Resolution Microscopy and Particle-Tracking Approaches for the Study of Vesicular Trafficking in Primary Neutrophils. . . . . . . . . . . . . . . . . Jennifer L. Johnson, Kersi Pestonjamasp, William B. Kiosses, and Sergio D. Catz An Approach to Monitor Exocytosis in White Adipocytes. . . . . . . . . . . . . . . . . . . . Ali M. Komai, Man Mohan Shrestha, Saliha Musovic, and Charlotta S. Olofsson Amperometry in Single Cells and Tissue . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Damien J. Keating Measurements of Exocytosis by Capacitance Recordings and Calcium Uncaging in Mouse Adrenal Chromaffin Cells. . . . . . . . . . . . . . . . . . . . . . Se´bastien Houy, Joana S. Martins, Ralf Mohrmann, and Jakob Balslev Sørensen Retention Using Selective Hooks-Synchronized Secretion to Measure Local Exocytosis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Gaelle Boncompain, Lou Fourriere, Nelly Gareil, and Franck Perez Combining Single Molecule Super-Resolution Imaging Techniques to Unravel the Nanoscale Organization of the Presynapse . . . . . . . . . . . . . . . . . . . Christopher Small, Ramon Martı´nez-Ma´rmol, Rumelo Amor Frederic A. Meunier, and Merja Joensuu Induction of Ca2+-Dependent Exocytotic Processes by Laser Ablation of Endothelial Cells . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Arsila P. K. Ashraf, Sophia N. Koerdt, Nikita Raj, and Volker Gerke Transmission Electron Microscopy and Tomography on Plasma Membrane Sheets to Study Secretory Docking . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Franck Delavoie, Cathy Royer, Ste´phane Gasman Nicolas Vitale, and Sylvette Chasserot-Golaz Spatial and Temporal Aspects of Exocytosis Studied on the Isolated Plasma Membranes . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Ira Milosevic

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

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Contributors KARINA NOEL ALTAMIRANO • Instituto de Histologı´a y Embriologı´a de Mendoza (IHEM) “Dr. Mario H. Burgos”. CONICET. Facultad de Ciencias Me´dicas, Universidad Nacional de Cuyo, Mendoza, Argentina RUMELO AMOR • Queensland Brain Institute, The University of Queensland, Brisbane, QLD, Australia ARSILA P. K. ASHRAF • Institute of Medical Biochemistry, Centre for Molecular Biology of Inflammation, University of Mu¨nster, Mu¨nster, Germany ASRAL W. ASNAWI • Department of Biomedical Science, Centre for Membrane Interactions and Dynamics, University of Sheffield, Sheffield, UK; Faculty of Medicine and Health Sciences, Universiti Sains Islam Malaysia, Negeri Sembilan, Malaysia NURIT P. AZOUZ • Department of Cell and Developmental Biology, Sackler Faculty of Medicine, Tel Aviv University, Tel Aviv, Israel; Department of Pediatrics, Cincinnati Children Hospital Medical Center, University of Cincinnati College of Medicine, Cincinnati, OH, USA MARIE-FRANCE BADER • Centre National de la Recherche Scientifique, Universite´ de Strasbourg, Institut des Neurosciences Cellulaires et Inte´gratives, Strasbourg, France SILVIA ALEJANDRA BELMONTE • Instituto de Histologı´a y Embriologı´a de Mendoza (IHEM) “Dr. Mario H. Burgos”. CONICET. Facultad de Ciencias Me´dicas, Universidad Nacional de Cuyo, Mendoza, Argentina GAELLE BONCOMPAIN • Dynamics of Intracellular Organization Laboratory, Institut Curie, PSL Research University, Sorbonne Universite´, Centre National de la Recherche Scientifique, UMR 144, Paris, France EMMANUEL BOUCROT • Institute of Structural and Molecular Biology, University College London, London, UK; Institute of Structural and Molecular Biology, Birkbeck College, London, UK DAVID A. CARDOSO • Cell Signalling Unit, Children’s Medical Research Institute, Faculty of Medicine and Health, The University of Sydney, Westmead, NSW, Australia SERGIO D. CATZ • Department of Molecular Medicine, The Scripps Research Institute, La Jolla, CA, USA MARA CERIDONO • Centre National de la Recherche Scientifique, Universite´ de Strasbourg, Institut des Neurosciences Cellulaires et Inte´gratives, Strasbourg, France SYLVETTE CHASSEROT-GOLAZ • Centre National de la Recherche Scientifique, Universite´ de Strasbourg, Institut des Neurosciences Cellulaires et Inte´gratives, Strasbourg, France NGOC CHAU • Cell Signalling Unit, Children’s Medical Research Institute, Faculty of Medicine and Health, The University of Sydney, Westmead, NSW, Australia MICHAEL A. COUSIN • Centre for Discovery Brain Sciences, Hugh Robson Building, George Square, University of Edinburgh, Edinburgh, Scotland, UK; Muir Maxwell Epilepsy Centre, Hugh Robson Building, George Square, University of Edinburgh, Edinburgh, Scotland, UK; Simons Initiative for the Developing Brain, Hugh Robson Building, George Square, University of Edinburgh, Edinburgh, Scotland, UK FRANCK DELAVOIE • Centre National de la Recherche Scientifique, Laboratoire de Biologie Mole´culaire Eucaryote, Centre de Biologie Inte´grative, Universite´ de Toulouse, Toulouse, France

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Contributors

ESTELLE DRANSART • Institut Curie, PSL Research University, Cellular and Chemical Biology Unit, Endocytic Trafficking and Intracellular Delivery Team, U1143 INSERM, UMR3666 CNRS, 26 rue d’Ulm, Paris Cedex 05, France LOU FOURRIERE • Dynamics of Intracellular Organization Laboratory, Institut Curie, PSL Research University, Sorbonne Universite´, Centre National de la Recherche Scientifique, UMR 144, Paris, France NELLY GAREIL • Dynamics of Intracellular Organization Laboratory, Institut Curie, PSL Research University, Sorbonne Universite´, Centre National de la Recherche Scientifique, UMR 144, Paris, France STE´PHANE GASMAN • Centre National de la Recherche Scientifique, Universite´ de Strasbourg, Institut des Neurosciences Cellulaires et Inte´gratives, Strasbourg, France VOLKER GERKE • Institute of Medical Biochemistry, Centre for Molecular Biology of Inflammation, University of Mu¨nster, Mu¨nster, Germany DAVID E. GORDON • Department of Cellular and Molecular Pharmacology, University of California San Francisco, San Francisco, CA, USA CLAUDIA HINZE • Institute of Structural and Molecular Biology, University College London, London, UK; Division of Infection and Immunity, Institute of Immunity and Transplantation, University College London, London, UK SE´BASTIEN HOUY • Department of Neuroscience, University of Copenhagen, Copenhagen N, Denmark MERJA JOENSUU • Clem Jones Centre for Ageing Dementia Research, Queensland Brain Institute, The University of Queensland, Brisbane, QLD, Australia LUDGER JOHANNES • Institut Curie, PSL Research University, Cellular and Chemical Biology Unit, Endocytic Trafficking and Intracellular Delivery Team, U1143 INSERM, UMR3666 CNRS, 26 rue d’Ulm, Paris Cedex 05, France JENNIFER L. JOHNSON • Department of Molecular Medicine, The Scripps Research Institute, La Jolla, CA, USA JERNEJ JORGACˇEVSKI • Laboratory of Neuroendocrinology—Molecular Cell Physiology, Institute of Pathophysiology, Faculty of Medicine, University of Ljubljana, Ljubljana, Slovenia; Celica BIOMEDICAL, Ljubljana, Slovenia MICHAEL A. KALWAT • Lilly Diabetes Center of Excellence, Indiana Biosciences Research Institute, Indianapolis, IN, USA DAMIEN J. KEATING • Flinders Health and Medical Research Institute, Flinders University, Adelaide, SA, Australia WILLIAM B. KIOSSES • Department of Vascular and Cell Biology, La Jolla Institute for Immunology, La Jolla, CA, USA OFIR KLEIN • Department of Cell and Developmental Biology, Sackler Faculty of Medicine, Tel Aviv University, Tel Aviv, Israel; Department of Molecular Genetics, Weizmann Institute of Science, Rehovot, Israel SOPHIA N. KOERDT • Institute of Medical Biochemistry, Centre for Molecular Biology of Inflammation, University of Mu¨nster, Mu¨nster, Germany ALI M. KOMAI • Department of Physiology/Metabolic Physiology, Institute of Neuroscience and Physiology, The Sahlgrenska Academy, University of Gothenburg, Go¨teborg, Sweden RAMON MARTI´NEZ-MA´RMOL • Clem Jones Centre for Ageing Dementia Research, Queensland Brain Institute, The University of Queensland, Brisbane, QLD, Australia JOANA S. MARTINS • Department of Neuroscience, University of Copenhagen, Copenhagen N, Denmark

Contributors

xi

KIERAN MCGOURTY • Institute of Structural and Molecular Biology, University College London, London, UK; Department of Chemical Sciences, Bernal Institute and Health Research Institute, University of Limerick, IRL, Limerick, Ireland FREDERIC A. MEUNIER • Clem Jones Centre for Ageing Dementia Research, Brisbane, QLD, Australia; Queensland Brain Institute, The University of Queensland, Brisbane, QLD, Australia IRA MILOSEVIC • European Neuroscience Institute (ENI), A Joint Initiative of the University Medical Center, Go¨ttingen and the Max Planck Society, Go¨ttingen, Germany; Wellcome Centre for Human Genetics, Nuffield Department of Medicine, NIHR Oxford Biomedical Research Centre, University of Oxford, Oxford, UK RALF MOHRMANN • Institute for Physiology, Otto-von-Guericke University, Magdeburg, Germany ALFONSINA MORALES • Instituto de Histologı´a y Embriologı´a de Mendoza (IHEM) “Dr. Mario H. Burgos”. CONICET. Facultad de Ciencias Me´dicas, Universidad Nacional de Cuyo, Mendoza, Argentina SALIHA MUSOVIC • Department of Physiology/Metabolic Physiology, Institute of Neuroscience and Physiology, The Sahlgrenska Academy, University of Gothenburg, Go¨teborg, Sweden CHARLOTTA S. OLOFSSON • Department of Physiology/Metabolic Physiology, Institute of Neuroscience and Physiology, The Sahlgrenska Academy, University of Gothenburg, Go¨teborg, Sweden STE´PHANE ORY • Centre National de la Recherche Scientifique, Universite´ de Strasbourg, Institut des Neurosciences Cellulaires et Inte´gratives, Strasbourg, France ANDREW A. PEDEN • Department of Biomedical Science, Centre for Membrane Interactions and Dynamics, University of Sheffield, Sheffield, UK FRANCK PEREZ • Dynamics of Intracellular Organization Laboratory, Institut Curie, PSL Research University, Sorbonne Universite´, Centre National de la Recherche Scientifique, UMR 144, Paris, France KERSI PESTONJAMASP • Department of Molecular Medicine, The Scripps Research Institute, La Jolla, CA, USA MAJA POTOKAR • Laboratory of Neuroendocrinology—Molecular Cell Physiology, Institute of Pathophysiology, Faculty of Medicine, University of Ljubljana, Ljubljana, Slovenia; Celica BIOMEDICAL, Ljubljana, Slovenia JULIETTE RAHERINDRATSARA • Centre National de la Recherche Scientifique, Universite´ de Strasbourg, Institut des Neurosciences Cellulaires et Inte´gratives, Strasbourg, France NIKITA RAJ • Institute of Medical Biochemistry, Centre for Molecular Biology of Inflammation, University of Mu¨nster, Mu¨nster, Germany PHILLIP J. ROBINSON • Cell Signalling Unit, Children’s Medical Research Institute, Faculty of Medicine and Health, The University of Sydney, Westmead, NSW, Australia CATHY ROYER • Plateforme Imagerie In Vitro, Neuropoˆle de Strasbourg, Strasbourg, France RONIT SAGI-EISENBERG • Department of Cell and Developmental Biology, Sackler Faculty of Medicine, Tel Aviv University, Tel Aviv, Israel LAURA SALAVESSA • Group Intracellular Trafficking and Tissue Homeostasis. Unite´ de Pathoge´nie Microbienne Mole´culaire, Institut Pasteur, Paris, France; U1202, INSERM, Paris, France; Universite´ Paris Sud, Paris-Saclay University, Orsay, France NATHALIE SAUVONNET • Group Intracellular Trafficking and Tissue Homeostasis. Unite´ de Pathoge´nie Microbienne Mole´culaire, Institut Pasteur, Paris, France; U1202, INSERM, Paris, France

xii

Contributors

MASSIULLAH SHAFAQ-ZADAH • Institut Curie, PSL Research University, Cellular and Chemical Biology Unit, Endocytic Trafficking and Intracellular Delivery Team, U1143 INSERM, UMR3666 CNRS, 26 rue d’Ulm, Paris Cedex 05, France MAN MOHAN SHRESTHA • Department of Physiology/Metabolic Physiology, Institute of Neuroscience and Physiology, The Sahlgrenska Academy, University of Gothenburg, Go¨teborg, Sweden AMBER S. SHUN-SHION • Department of Biomedical Science, Centre for Membrane Interactions and Dynamics, University of Sheffield, Sheffield, UK CHRISTOPHER SMALL • Clem Jones Centre for Ageing Dementia Research, Brisbane, QLD, Australia; Queensland Brain Institute, The University of Queensland, Brisbane, QLD, Australia KAREN J. SMILLIE • Centre for Discovery Brain Sciences, Hugh Robson Building, George Square, University of Edinburgh, Edinburgh, Scotland, UK; Muir Maxwell Epilepsy Centre, Hugh Robson Building, George Square, University of Edinburgh, Edinburgh, Scotland, UK JAKOB BALSLEV SØRENSEN • Department of Neuroscience, University of Copenhagen, Copenhagen N, Denmark LAILA SUHAIMAN • Instituto Interdisciplinario de Ciencias Ba´sicas (ICB) CONICET. Facultad de Ciencias Exactas y Naturales, Universidad Nacional de Cuyo, Mendoza, Argentina EMELINE TANGUY • Centre National de la Recherche Scientifique, Universite´ de Strasbourg, Institut des Neurosciences Cellulaires et Inte´gratives, Strasbourg, France TAMOU THAHOULY • Centre National de la Recherche Scientifique, Universite´ de Strasbourg, Institut des Neurosciences Cellulaires et Inte´gratives, Strasbourg, France NICOLAS VITALE • Centre National de la Recherche Scientifique, Universite´ de Strasbourg, Institut des Neurosciences Cellulaires et Inte´gratives, Strasbourg, France ROBERT ZOREC • Laboratory of Neuroendocrinology—Molecular Cell Physiology, Institute of Pathophysiology, Faculty of Medicine, University of Ljubljana, Ljubljana, Slovenia; Celica BIOMEDICAL, Ljubljana, Slovenia

Part I Endocytosis

Chapter 1 Stoichiometry of Receptors at the Plasma Membrane During Their Endocytosis Using Total Internal Reflection Fluorescent (TIRF) Microscopy Live Imaging and Single-Molecule Tracking Laura Salavessa and Nathalie Sauvonnet Abstract Determination of protein stoichiometry in living cells is key to understanding basic biological processes. This is particularly important for receptor-mediated endocytosis, a highly regulated mechanism that requires the sequential assembly of numerous factors. Here, we describe a quantitative approach to analyze receptor clustering dynamics at the plasma membrane. Our workflow combines TIRF live imaging of a CRISPR-Cas9-edited cell line expressing a GFP-tagged receptor in a physiological relevant environment, a calibration technique for single-molecule analysis of GFP, and detection and tracking with an open-source software. This method allows to determine the number of receptor molecules at the plasma membrane in real time. Key words Single-molecule tracking (SPT), TIRF, Endocytosis, Photobleaching, Receptor, Dynamics, Plasma membrane, Quantitative biology

1

Introduction Many proteins are part of multimeric complexes, often containing different number and type of subunits, and oligomerize as part of their catalytic activity or function. Such proteins play a critical role in a wide variety of cellular events, which is particularly notable in membrane proteins that drive ion and protein transport and homeostasis, signal transduction, endocytosis, and secretion. Therefore, studying their stoichiometry and dynamics, under native conditions in living cells, is key to better understand their function and the cellular mechanisms in which they participate. In this introduction, we will briefly present some examples reported in the literature, describing the link between protein stoichiometry and function.

Florence Niedergang, Nicolas Vitale and Ste´phane Gasman (eds.), Exocytosis and Endocytosis: Methods and Protocols, Methods in Molecular Biology, vol. 2233, https://doi.org/10.1007/978-1-0716-1044-2_1, © Springer Science+Business Media, LLC, part of Springer Nature 2021

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One of the first well-studied examples is the Epidermal Growth Factor Receptor (EGFR). Although largely accepted that ligandinduced dimerization of EGFR is essential for signal transduction, some studies showed that EGFR activation is also associated with the formation of high-order oligomers that efficiently propagate the signal [1]. Indeed, several studies analyzing EGFR stoichiometry using Fo¨rster Resonance Energy Transfer (FRET), Fluorophore Localization Imaging with Photobleaching (FLImP), or biochemical techniques like cross-linking experiments [2] revealed that formation of at least receptor tetramers was necessary for higher tyrosine kinase activity [1, 3]. Moreover, mutations blocking EGFR multimerization have been shown to reduce its autophosphorylation and the activation of downstream signaling partners [3, 4]. Altogether, these findings show that EGF signaling is regulated by receptor clustering, which is of particular interest since mutations in EGFR that lead to enhanced signaling are frequently associated with cancer. Another example for which receptor stoichiometry has been linked to its function is CC-chemokine receptor 5 (CCR5), a G Protein-Coupled Receptor (GPCR) and the main coreceptor for human immunodeficiency virus type 1 (HIV-1). Different affinities of the virus for CCR5 have been proposed to regulate HIV virulence, resistance to inhibitors, and cellular tropisms. In particular, using bimolecular fluorescence complementation (BiFC) and FRET assays, it was shown that various CCR5 populations are present at the cell surface and that these exist in a variety of conformations due to different dimerization interfaces. Furthermore, it was shown that HIV-1 preferentially recognizes CCR5 monomers as opposed to dimers [5, 6], in agreement with findings that anti-CCR5 monoclonal antibodies that induce receptor dimerization prevent HIV-1 infection [7]. Therefore, the oligomerization state of CCR5 seems to regulate HIV interaction, uptake, and infection. It should be noted that, for most of the stoichiometry analysis summarized above, EGFR and CCR5 were ectopically expressed in cells, which might result in a biased analysis due to the nonphysiological expression of the receptors. Another mechanism for which protein stoichiometry has been shown to be determinant is endocytosis. The highly characterized clathrin-mediated uptake is a process involving many factors [8], although the orchestration of these factors and their organization is not fully understood. Recent advances in gene-editing tools and single-molecule fluorescence imaging techniques enabled the characterization of protein oligomerization and complex assembly at the plasma membrane in living cells. For instance, in clathrinmediated endocytosis, pits initiate by coordinated arrival of one clathrin triskelion and two membrane-bound AP2 proteins that stabilize the complex, increasing its residence time at the membrane

Single-Molecule Analysis in Real Time at the Plasma Membrane

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and allowing further addition of more clathrin molecules [9]. In mammals, the last step of vesicle fission requires the action of the large GTPase dynamin. Recently, several groups showed that approximately 26 molecules of dynamin are recruited to the neck of the clathrin vesicle [10, 11], which is sufficient to form a short helix that encircles the neck of the vesicle once and bridges two dynamin molecules together, thus activating the GTPase and driving scission [10]. Altogether, these studies demonstrate that understanding protein stoichiometry and assembly is necessary to provide new insights, not only in its physiological function but also in its biogenesis, its targeting by pathogens, and its deregulation in disease. In this chapter, we will describe our method to infer in realtime the number of proteins at the plasma membrane, using an endogenously expressed gene-edited receptor coupled to a fluorescent tag and single-molecule analysis. To quantify the number of molecules at the plasma membrane, we take advantage of the high signal-to-noise ratio provided by TIRF microscopy. Our receptor model is the γ chain of the interleukin-2 receptor (IL-2Rγ), a critical signaling receptor for immune function and a fine example of a clathrin- and caveolin-independent endocytic cargo [12, 13]. The receptor was found to accumulate at the base of membrane protrusions, suggesting this could constitute the initial step of endocytic pit formation [14]. This endocytic process does not depend on any coat protein, thus raising the question as to whether local receptor clustering may initiate membrane bending. To test this hypothesis, we developed a workflow to analyze IL-2Rγ stoichiometry during endocytosis in living T lymphocytes. Our protocol combines TIRF live imaging of CRISPR-Cas9edited cells that endogenously express the receptor of interest coupled to eGFP, with a calibration method for single-molecule analysis using purified eGFP to determine the unitary bleaching step of the fluorophore. The intensity of a single eGFP molecule corresponds to the loss of fluorescence signal observed in one photobleaching step. This intensity standard (unitary bleaching step) is then used to infer the number of receptor molecules, in live-cell TIRF images, that are tracked with a plug-in implemented in the open-source software Icy (Fig. 1). Altogether, this method reports the number of receptor molecules at the plasma membrane during endocytosis, in real time.

2

Materials

2.1 Coverslips and Cell Preparation

1. CRISPR-Cas9-edited cell line, stably expressing the receptor of interest with a fluorophore like eGFP, under its own promoter (see Note 1). In our particular example, we use a T-cell line

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Fig. 1 Workflow for single-molecule analysis of plasma membrane receptor stoichiometry. (a) Schematic representation of cell seeding onto poly-L-lysine-coated coverslips and of the O-ring coverslip holder used during imaging. (b) TIRF imaging allows observation of molecules close to the coverslip as purified eGFP and plasma membrane receptors. (c) Calibration for eGFP single-molecule analysis relies on photobleaching of eGFP molecules and detection of bleaching steps upon high-frequency acquisition. (d) Detection and tracking of surface receptors are done using the eTrack plug-in in Icy software

(Kit225 cells [15]) expressing the γ chain of the IL-2R tagged with eGFP (GFP-IL-2Rγ). 2. Roswell Park Memorial Institute (RPMI) 1640 medium containing L-glutamine and phenol red, and supplemented with 10% fetal bovine serum (heat inactivated) and 1 nM IL-2, for cell culture at 37
C. 3. Round 25 mm diameter glass No. 01 (0.13–0.17 mm thickness) (see Note 2). 4. Ethanol 70%. 5. Acetone. 6. Ultrasonic cleaning bath (220 V, 50/60 Hz). 7. 6-well plate. 8. ddH2O. 9. Poly-L-lysine 0.1% (w/v).

coverslips

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2.2

TIRF Imaging

7

1. O-ring Attofluor cell chamber (coverslip holder). 2. Purified eGFP (1 mg/mL). 3. PBS 1. 4. Cell imaging medium consisting of RPMI 1640 medium without phenol red, supplemented with 5% fetal bovine (heat inactivated). 5. Immersion oil with a refractive index of 1.518, suited for 37
C imaging.

2.2.1 Live-Image Acquisition Is Performed with an Inverted Confocal Microscope, LSM 780 Elyra PS.1 Equipped with an EMCCD Camera. The Setup Used Includes the Following Components

1. LSM 780 Elyra PS.1 inverted microscope, equipped with a motorized XY stage, an environmental chamber with temperature control, and a focus control by definite focus. 2. A 488 nm (100 mW) HR—solid laser line for eGFP excitation. 3. A filter cube set with a band-pass (BP) 495–575 + low-pass (LP) 750 filter for eGFP detection. 4. An alpha Pin Apo 100/1.46 numerical aperture (NA) oil objective. 5. An EMCCD 887 1K camera. 6. ZEN software.

2.3

Image Analysis

Detection and tracking of purified eGFP and of membrane receptor are performed with Icy software (www.icy.bioimageanalysis.org) [16] and then further analyzed using Microsoft Excel. 1. Icy software version 2.0.2.0, containing the plug-ins: Spot Detector, ROI Intensity Evolution, eTrack, and Intensity Track Processor. 2. Microsoft Excel.

3

Methods

3.1 Coverslips and Cell Preparation

Glass coverslips should be washed prior to use. A washing step with ethanol and acetone, followed by an ultrasonic cleaning bath, enables removal of any oil/grease films from the surface of the glass and ensures optimal adherence of cells with minimal background. Afterwards, coverslips are coated with poly-L-lysine when working with suspension cells like T cells (Fig. 1a). This poly-Llysine step is unnecessary when using adherent cells. 1. Clean coverslips by immersing glass coverslips in 70% ethanol for 20 min, then rinse in ddH2O three times, and sonicate in an ultrasonic cleaning bath for 5 min. After, the process is repeated by immersing coverslips in acetone instead of ethanol.

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2. Coat washed coverslips by placing them in a 6-well plate with poly-L-lysine diluted in ddH2O in a ratio of 1:10 for 20 min. Coverslips are then washed twice with ddH2O, placed separately in a coverslip rack, and allowed to dry completely. Coated coverslips can be kept at 4
C, in a sterile sealed container, for up to 3 months. 3. Wash Kit225 GFP-IL-2Rγ once with medium without serum, and then seed them in the same medium at approximately 1.5 million cells per coverslip, in a 6-well plate (see Note 3). The plate is centrifuged at 300  g for 2 min to promote cell adherence. Cells can then be kept in regular RPMI medium, at 37
C with 5% CO2, until imaging. 3.2

TIRF Imaging

In order to calibrate the imaging system for the analysis of eGFP fluorescence at single-molecule level, purified eGFP should be coated onto coverslips at low density (Fig. 2a) and imaged at high acquisition frequency (see Note 4). This calibration step is essential and needs to be performed for each experiment, using the same parameters (TIRF angle, laser power, exposure time and gain) as for the live-cell imaging. Therefore, these parameters should be first set up on a live-cell sample in order to obtain an evanescent wave and avoid, as much possible, photobleaching of the eGFP-tagged receptor (see Note 5). Importantly, acquisition of both purified eGFP and living cells should be done under a uniform field of illumination (Fig. 1b). 1. Preheat at 37
C the microscope chamber and coverslip holder prior imaging to avoid focus problems. 2. Prepare the purified eGFP by diluting it with PBS 1 to 1:40000 in 1 mL (eGFP 25 ng/mL final concentration), and add it to a glass coverslip placed on a pre-warmed coverslip holder and incubate for 30 s at 37
C, in the microscope chamber. After, the eGFP-coated coverslip is washed thoroughly five times and imaged in PBS. 3. Set the TIRF acquisitions using the 100/1.46 NA oil objective and ZEN software, with the 488 nm channel. The settings chosen, including exposure time, gain, laser power, and TIRF angle, must be identical to the ones used during cell imaging. As an example, the settings used were as follows: exposure time of 150 ms, gain of 300, laser power of 1%, and TIRF angle of 64.4
. The eGFP sample is imaged at high frequency of 20 Hz (1 frame/50 ms) and recorded for a total of 300 frames. 4. Live-image the surface receptors by placing a coverslip seeded with cells on a pre-warmed coverslip holder, and carefully add 1 mL of imaging medium to it. 5. Image the cells at 37
C using the same settings as for purified eGFP imaging, with the exception of acquisition rate, which for

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Fig. 2 Calibration for eGFP single-molecule photobleaching detection. (a) Example images of a correct density of eGFP seeded onto coverslips and an incorrect, overly dense, eGFP seeding that should be avoided. (b) Detection of eGFP spots on the first frame of the eGFP high-frequency acquisition movie using the Spot Detector plug-in in Icy. Highlighted in orange are the parameters to detect bright spots over dark background, scale and sensitivity for spot detection, and the tab Output for export of spots as ROIs (step 2 of Subheading 3.3). (c) Conversion of ROIs into circles with a fixed radius of two pixels from their barycenter (highlighted), copy of these ROIs onto all frames of the eGFP movie (highlighted), and deletion of overlapping and border spots (highlighted) (step 3 of Subheading 3.3). (d) Detection of eGFP fluorescence intensity along time with the ROI Intensity Evolution plug-in. All spots are selected (highlighted), and their intensities are saved as an Excel file (step 4 of Subheading 3.3). (e) Example plots of eGFP fluorescence intensity along time, with an exponential decrease in intensity where no discrete bleaching step is observable (left), and one plot with a clear bleaching step (right). In plots with observable bleaching steps, the mean fluorescence intensity before (m1) and after (m2) the bleaching step is calculated, and their difference represents the photobleaching of one eGFP molecule (step 5 of Subheading 3.3). (f) The relative frequency of the unitary bleaching steps of several eGFP spots follows a Gaussian distribution. The mean of the unitary bleaching step, here with a value of 400 a. u., is later used to calculate the number of receptor molecules in endocytic tracks (step 6 of Subheading 3.3)

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this assay was set to 1 Hz (1 frame/s). Cells are imaged for a total of 150 frames, and an average of 12–15 cells are imaged per condition per experiment. 3.3 Calibration for eGFP Single-Molecule Photobleaching Detection

In order to calculate the number of receptor molecules present in an endocytic pit, the unitary bleaching step of the fluorophore, in this case eGFP, has to be determined using the fast acquisition movies of purified eGFP (Fig. 1c). Icy software is used to obtain the fluorescence intensity along time of the imaged eGFP. During this step, eGFP spots are automatically detected according to their approximate pixel size (scale) and brightness (sensitivity) using a wavelet-based algorithm implemented in the plug-in Spot Detector [16]. The ROIs of the detected spots are exported, and their sizes are uniformized by converting the ROIs into circles with a fixed radius from their barycenter position. Later, the intensity of GFP-IL-2Rγ spots will be acquired using the same ROI size as for eGFP (see Note 6). By plotting the fluorescence intensity of each eGFP spot along time, it is possible to observe the bleaching step of one fluorophore and determine its intensity when discrete bleaching steps are observed (see Note 7). 1. Load the purified eGFP movie into Icy, and draw and crop a square ROI with a homogeneous illumination and a low eGFP density. 2. Detect eGFP spots by extracting the first frame of the movie, and use it as the input sequence for detection of spots with Spot Detector, within the tab Detection & Tracking (see Note 8). The plug-in is set to detect bright spots over dark background, in its Detector tab, and the scale and sensitivity of detection are chosen according to the spot size and brightness (see Note 9). In our case, a scale of 2 (objects of approximately 3–6 pixels size) and sensitivity of 70 are regularly used. In the Output tab of the plug-in, the Export to ROI is ticked (Fig. 2b). 3. Select and convert the exported ROIs into circles with a fixed size from their barycenter position in the section Conversion in the tab Region of Interest. A radius of 2 pixels was used for the conversion. All spots (ROIs) are then selected from the ROI panel and copied into the eGFP movie, making sure the copied ROIs are applied to all frames (T ¼ ALL in the ROI panel). Spots that are placed on the borders of the image, and overlapping spots are suppressed (Fig. 2c). 4. Plot the intensity of the eGFP spots over time using the ROI Intensity Evolution plug-in. The eGFP movie is selected as the sequence to analyze and all ROIs are selected. Once the intensity plots of the ROIs are displayed, the results can be saved as an XLS file (Fig. 2d).

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5. Further analyze eGFP intensities using Microsoft Excel (see Note 10). The exported data shows the mean intensity of eGFP per frame, with each column corresponding to one eGFP detected spot. Individual columns are selected, and their intensity along time is plotted. Spots that do not show clear bleaching steps are deleted (see Note 7 and Fig. 2e). Then, the mean fluorescence intensity before and after the bleaching step is calculated, and their difference corresponds to the unitary bleaching step of one molecule of eGFP (Fig. 2e). 6. Calculate the unitary bleaching steps for approximately 100 spots. If plotted, the bleaching step intensity of these spots should follow a Gaussian frequency distribution (Fig. 2f). If otherwise, it might indicate an excessive density and aggregation of eGFP. The mean of the unitary bleaching steps (UeGFP) corresponds to the intensity value of one molecule of eGFP and is later used to infer the number of GFP-IL2Rγ molecules. 3.4 Single-Molecule Analysis of Surface Receptors

When imaging receptors in living cells, the dynamics of the process have to be preserved, which prevents direct counting of molecules by photobleaching. In this case, the fluorescence intensity of the eGFP-tagged receptor can be used and divided by the unitary bleaching step of eGFP (previously determined) to infer the number of molecules in a complex. The eTrack plug-in implemented in Icy is used for detection of surface receptors on live-imaging movies (Fig. 1d). This plug-in was developed for the detection of putative endocytic tracks (see Note 11) and uses Spot Detector to identify spots that are significantly brighter than the background, generating an elevation map of their fluorescence for the whole duration of the movie. In this elevation map, putative endocytic sites appear as high-intensity Gaussian peaks due to the multiple spot detections that are summed along the frames. In addition, eTrack relies on the xy confinement of endocytic tracks over time to precisely determine the position of putative endocytic sites and successfully reconstruct them even if some detections are missing. Not all parameters of the plug-in will be mentioned here, yet the default values are usually a good starting point. The reader can refer to Bertot et al. for a description of eTrack and example movies (Movie S1 and S2) [17]. 1. Load live-cell imaging movies into Icy, draw an ROI surrounding the cell (Fig. 3a), and use Spot Detector to determine the best parameters of scale and sensitivity of detection to be later introduced in the eTrack plug-in (see Note 9). These parameters may vary slightly among experiments but should be kept the same within the same experiment.

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Fig. 3 Single-molecule analysis of surface receptors. (a) Example of a TIRF movie with the ROI of one cell outlined (step 1 of Subheading 3.4). (b) eTrack panel with the parameters of threshold for spot detection, maximum gap closing, and minimum tube duration highlighted (step 2 of Subheading 3.4). (c) The resulting detected tracks will be shown on the movie (left), and their fluorescence intensity can be obtained by adding the Intensity Track Processor to the Track Manager panel (highlighted). All tracks are selected, the Time lag to extend tracks (highlighted) and the Diameter of the disk area in pixels (on the right, not shown) are defined, and results are exported to Excel (step 3 of Subheading 3.4). (d) GFP-IL-2Rγ at the cell surface with two tracks highlighted and example calculations to determine their stoichiometry based on their fluorescence intensity (MAX Imean), the background fluorescence of the cell (MIN Imin), and the mean of the unitary bleaching steps of eGFP (MEAN UeGFP) (step 4 of Subheading 3.4)

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2. Analyze live-cell imaging by launching the plugin eTrack in the tab Detection & Tracking. The live-cell imaging movie to be analyzed is selected as the input sequence, and the parameters previously chosen for spot detection are inserted, ensuring the plug-in is set to detect bright spots in dark background. In the advance parameters, it is possible to specify a variety of settings such as the minimum duration of the tracks (in frames) and the maximum number of missed detections (in frames) between real detections within a track (maximum gap between detections). In the case of GFP-IL-2Rγ, a scale 2 and threshold of 50 are generally used for detection, and the maximum gap closing and minimum tube duration are set to 4 and 15 frames, respectively. The results are exported to Excel, by defining a file name and location, and the plug-in is started (Fig. 3b). 3. Obtain the fluorescence intensity of these tracks using the TrackManager window (automatically open), in which all detected tracks can be found. To obtain the fluorescence intensity of these tracks, first all tracks must be selected (Edit > Select all tracks) and then the Intensity Track Processor is added to TrackManager. This new processor will show as a drop-down box named Intensity profile with two parameters: the Time lag to extended tracks defines the number of frames, before and after each track, for which fluorescence intensity will be additionally recorded, and the Averaging type defines the shape of the ROI, which is usually set to DISK, with a defined Diameter of the disk area in pixels. In our case, we do not extend the fluorescence recording of the tracks (Time lag to extended tracks ¼ 0), and we detect the fluorescence intensity of the tracks in a disk ROI with four pixels of diameter, matching the purified eGFP ROIs previously obtained. Next, the tracks are added to the processor by clicking in Put current selection in trackset A, and results are exported to Excel (Fig. 3c). 4. Obtain the number of molecules per track by performing a post analysis of the track fluorescent intensity with the two Excel files newly created. The first one contains information on the receptor tracks (from eTrack, step 2), such as xy position, duration of the track, initial and final frame, number of spots (real detections), mean spot size, and others. The second file contains data on the mean, maximum, and minimum fluorescence intensity of each track (from Track Manager, step 3). Importantly, the track numbers between the two files are corresponding; therefore, the fluorescence intensity of a track can be related with its duration, for example. Using the latter file, the background fluorescence of the cell is obtained by retrieving the minimum value of intensity (Excel function MIN) among all frames and tracks, in the Excel worksheet Channel 0—Min intensity (see Note 12). Then, in the

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worksheet Channel 0—Mean intensity, the maximum value of intensity (Excel function MAX), for each track, is retrieved, and the background value of intensity, previously obtained, is subtracted from it. This is the fluorescence intensity value used to calculate the number of molecules in a receptor track, by simply diving it by the mean of the unitary bleaching steps of eGFP. This calculation can be described as: N molecules ¼

MAX I mean  MIN I min MEAN U eGFP

Nmolecules: Number of receptor molecules in a track. MAX Imean: Maximum value of the mean intensity of the track. MIN Imin: Minimum value of the minimum intensity of all the tracks, corresponding to the cell’s fluorescence background and being a constant value used for all tracks within that cell. MEAN UeGFP: Mean of the unitary bleaching step values of purified eGFP (see step 6 of Subheading 3.3). An example of this calculation for two distinct tracks is given in Fig. 3d.

4

Notes 1. Single-molecule photobleaching experiments rely on bright and photostable fluorophores, with eGFP being the most commonly used. Expression of receptors fused to a fluorescent protein, protein tags as SNAP- or HALO-tags, or a fluorescent ligand or antibody can be used to visualize membrane-located proteins, such as receptors. However, the presence of endogenous non-labelled receptor and the degree and stoichiometry of labeling has to be accounted for in these situations. Therefore, genome editing methods are ideal to generate cell lines in which the fluorescent tag is expressed coupled to the endogenous gene of the receptor in both alleles, thus overcoming these issues. 2. Alternatively, glass bottom MatTek plates can be used although glass coverslips allow for a better cleaning procedure. 3. Cell density should be adjusted to at least 50% confluency according to cell type. T cells are particularly small hence a higher cell density is required to cover the coverslip. It is crucial to wash the cells and seed them in medium without serum. The presence of serum prevents proper adhesion of cells to the coverslip, regardless of the poly-L-lysine coating. 4. Coating of eGFP at low density (lower than 1–2 spots per μm2) is extremely important since single-molecule photobleaching counting is conditioned by the diffraction limit of light. At high

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densities of eGFP, single-molecule fluorescent spots will overlap or form aggregates, limiting the observation of single photobleaching steps. A dilution between 1:10000 and 1:100000 should be used for eGFP coating. The final density also varies greatly with the eGFP incubation time at 37
C and the number of washes. Additional washes can be done if, when starting acquisition, a slight excessive density of eGFP is seen at the microscope. Otherwise, if clustering of eGFP is noticeable, the best is to restart the preparation with an increased dilution. The reader can refer to Fig. 2a for an example of a correct and incorrect density of fluorophore coating. 5. Edited cells express endogenous amounts of the receptor coupled to one fluorophore, which usually results in a dim signal that is easily photobleached. Therefore, it is essential to choose a low laser power in order to reduce photobleaching of the tagged receptor in living cells. Additionally, low illumination intensity ensures a slow and observable stepwise photobleaching of the purified fluorophore. To increase the signal obtained, the exposure time and gain can be adjusted, taking into account that a higher exposure time will also lead to photobleaching, and a higher gain will increase background. A compromise between these parameters ensures the acquisition of suitable data for analysis. To achieve total internal reflection, the incident light has to arise from the objective at a greater angle than the critical angle for the coverslip-medium interface. In our experience, this corresponds to a TIRF angle between 62
and 65
. For a detailed description of TIRF imaging, refer to Yildiz et al. [18]. 6. The optimal radius for detection of spots should be determined so that the eGFP spots and the receptor spots fit within it yet avoiding overlapping with neighboring spots. 7. Discrete bleaching steps are seen when there is a clear abrupt decrease in fluorescence intensity of the eGFP as seen in the graph on the right in Fig. 2e. If eGFP is aggregated, multiple fluorophores will be clustered in one diffraction limited spot, and the fluorescence intensity will decrease exponentially (Fig. 2e graph on the left); hence, discrete bleaching steps will not be observed. 8. Alternatively, if the eGFP signal is dim, the initial 10 frames can be extracted and converted into a z-stack, and the maximum intensity projection is done. This might help with obtaining a brighter signal from the eGFP spots present during the first frames and thus a better detection with Spot Detector. 9. Different scales and sensitivities should be previously tested, according to the pixel size and brightness of the object of interest. For this optimization, the option Display binary

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image in the Detector tab of Spot Detector is of great use, allowing a clear visualization of the resulting ROIs. These parameters might vary between experiments, particularly if different conditions are being studied as, for instance, while investigating the effect of a drug on receptor endocytosis. 10. There are several “step detection” algorithms, implemented in MATLAB or other softwares, that can be used or modified to help in fast-automatic detection of single-molecule photobleaching steps, reducing the time of analysis [19, 20]. 11. eTrack relies on the xy confinement of endocytic tracks over time, which means that tracks that are detected for the minimum number of frames set in the plug-in (minimum duration of the tracks) but that diffuse afterwards will stop being tracked and, consequently, present short durations. These tracks are not endocytic tracks since tracking was interrupted due to diffusion instead of pit invagination and exit from the TIRF plane, which occurs upon endocytosis. Therefore, the classification of tracks obtained with eTrack has to be carefully done and depends on the purpose of the study. Alternatively, other tracking methods that consider xy displacement can be used. 12. The background fluorescence of the cell can also be obtained by extending the fluorescence detection of the receptor tracks, using the parameter Time lag to extended tracks in the Intensity Track Processor. This method leads to similar values of background as the determination of the minimum fluorescence intensity among all tracks. It has, however, the disadvantage that it will produce an extended fluorescence detection that might be contaminated by the fluorescence intensity of neighboring or diffusive spots that are in close proximity to the detected track. This extended fluorescence detection will always have to be considered and, preferentially, removed from further analysis.

Acknowledgments First, we would like to thank Dr. Alexandre Grassart for training us with the technologies of CRISPR/Cas9 gene edition and singlemolecule analysis and for all helpful discussion. We thank the PBI (Imagopole) platform of Institut Pasteur for microscope maintenance and technical help. LS is part of the Pasteur Paris University (PPU) International PhD Program and has received funding from the European Union’s Horizon 2020 research and innovation program under the Marie Sklodowska-Curie grant agreement No 665807.

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References 1. Clayton AHA, Walker F, Orchard SG et al (2005) Ligand-induced dimer-tetramer transition during the activation of the cell surface epidermal growth factor receptor-A multidimensional microscopy analysis. J Biol Chem 280:30392–30399 2. Liang SI, van Lengerich B, Eichel K et al (2018) Phosphorylated EGFR dimers are not sufficient to activate Ras. Cell Rep 22:2593–2600 3. Needham SR, Roberts SK, Arkhipov A et al (2016) EGFR oligomerization organizes kinase-active dimers into competent signalling platforms. Nat Commun 7:13307 4. Huang Y, Bharill S, Karandur D et al (2016) Molecular basis for multimerization in the activation of the epidermal growth factor receptor. Elife 5:e14107 5. Nakano Y, Monde K, Terasawa H et al (2014) Preferential recognition of monomeric CCR5 expressed in cultured cells by the HIV-1 envelope glycoprotein gp120 for the entry of R5 HIV-1. Virology 452–453:117–124 6. Colin P, Zhou Z, Staropoli I et al (2018) CCR5 structural plasticity shapes HIV-1 phenotypic properties. PLoS Pathog 14:e1007432 7. Vila-Coro AJ, Mellado M, Martı´n de Ana A et al (2000) HIV-1 infection through the CCR5 receptor is blocked by receptor dimerization. Proc Natl Acad Sci U S A 97:3388–3393 8. Kaksonen M, Roux A (2018) Mechanisms of clathrin-mediated endocytosis. Nat Rev Mol Cell Biol 19:313–326 9. Cocucci E, Aguet F, Boulant S et al (2012) The first five seconds in the life of a Clathrin-coated pit. Cell 150:495–507 10. Grassart A, Cheng AT, Hong SH et al (2014) Actin and dynamin2 dynamics and interplay during clathrin-mediated endocytosis. J Cell Biol 205:721–735 11. Cocucci E, Gaudin R, Kirchhausen T (2014) Dynamin recruitment and membrane scission

at the neck of a clathrin-coated pit. Mol Biol Cell 25:3595–3609 12. Lamaze C, Dujeancourt A, Baba T et al (2001) Interleukin 2 receptors and detergent-resistant membrane domains define a Clathrinindependent endocytic pathway. Mol Cell 7:661–671 13. Gesbert F, Sauvonnet N, Dautry-Varsat A (2004) Clathrin-lndependent endocytosis and signalling of interleukin 2 receptors IL-2R endocytosis and signalling. Curr Top Microbiol Immunol 286:119–148 14. Basquin C, Trichet M, Vihinen H et al (2015) Membrane protrusion powers clathrinindependent endocytosis of interleukin-2receptor. EMBO J:1–15 15. Hori T, Uchiyama T, Tsudo M et al (1987) Establishment of an interleukin 2-dependent human T cell line from a patient with T cell chronic lymphocytic leukemia who is not infected with human T cell leukemia/lymphoma virus. Blood 70:1069–1072 16. de Chaumont F, Dallongeville S, Chenouard N et al (2012) Icy: an open bioimage informatics platform for extended reproducible research. Nat Methods 9:690–696 17. Bertot L, Grassart A, Lagache T et al (2018) Quantitative and statistical study of the dynamics of Clathrin-dependent and -independent endocytosis reveal a differential role of EndophilinA2. Cell Rep 22:1574–1588 18. Yildiz A, Vale RD (2015) Total internal reflection fluorescence microscopy. Cold Spring Harb Protoc 2015:pdb.top086348 19. Chen Y, Deffenbaugh NC, Anderson CT et al (2014) Molecular counting by photobleaching in protein complexes with many subunits: best practices and application to the cellulose synthesis complex. Mol Biol Cell 25:3630–3642 20. Tsekouras K, Custer TC, Jashnsaz H et al (2016) A novel method to accurately locate and count large numbers of steps by photobleaching. Mol Biol Cell 27:3601–3615

Chapter 2 Measuring Endocytosis During Proliferative Cell Quiescence Claudia Hinze, Kieran McGourty, and Emmanuel Boucrot Abstract Quiescence (also called “G0”) is the state in which cells have exited the cell cycle but are capable to reenter as required. Though poorly understood, it represents one of the most prevalent cell states across all life. Many biologically important cell types reside in quiescence including mature hepatocytes, endothelial cells, and dormant adult stem cells. Furthermore, the quiescence program occurs in both short- and long-term varieties, depending on the physiological environments. A barrier slowing our understanding of quiescence has been a scarcity of available in vitro model systems to allow for the exploration of key regulatory pathways, such as endocytosis. Endocytosis, the internalization of extracellular material into the cell, is a fundamental and highly regulated process that impacts many cell biological functions. Accordingly, we have developed an in vitro model of deep quiescence in hTERT-immortalized RPE1 cells, combining both longterm contact inhibition and mitogen removal, to measure endocytosis. In addition, we present an analytical approach employing automated high-throughput microscopy and image analysis that yields high-content data allowing for meaningful and statistically robust interpretation. Importantly, the methods presented herein provide a suitable platform that can be easily adapted to investigate other regulatory processes across the cell cycle. Key words Cell cycle, Cell quiescence, G0, Primary cells, hTERT-immortalized cells, Endocytosis, Clathrin-mediated endocytosis, Macropinocytosis, Fluid-phase uptake, Epidermal growth factor, Automated high-throughput microscopy, High-throughput image analysis

1

Introduction Proliferative quiescence (also known as the “G0” stage of the cell cycle) is the most common cell state among all cellular organisms and is defined as a temporary and reversible absence from proliferation [1]. This is different from senescence or terminal cell differentiation, which are irreversible exits from the cell cycle. Quiescent cells, such as most adult stem cells, are actively maintained in a quiescent state but can reenter the cell cycle when stimulated to rapidly expand and differentiate, enabling efficient tissue homeostasis. As such, quiescence is a default state that cells revert to when faced with a challenge to proliferation such as a lack of nutrients, mitogen signaling, or space (contact inhibition) [2–6]. It is also a

Florence Niedergang, Nicolas Vitale and Ste´phane Gasman (eds.), Exocytosis and Endocytosis: Methods and Protocols, Methods in Molecular Biology, vol. 2233, https://doi.org/10.1007/978-1-0716-1044-2_2, © Springer Science+Business Media, LLC, part of Springer Nature 2021

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mechanism for cells to preserve their function over a long period of time. Quiescent cells are defined by having a diploid (“2N”) genome [7], low levels of cell cycle markers (e.g., Ki67, nuclear Cyclin D1, or phosphorylated retinoblastoma protein), high levels of cyclin-dependent kinase inhibitors p21Cip1 and p27Kip1 [4, 8], elevated autophagy activity [9], and decreased levels and translation of mRNA [6, 10], as well as having a smaller cell size and an increased volume ratio of nucleus to cytoplasm [4, 5, 11]. The traditional understanding of cell quiescence was that the cell was completely dormant and stagnant (e.g., the “sleeping beauty” state in Baker’s yeast) [7]. As more research has been carried out on proliferative quiescence, new evidence showed that this cell state is dynamic and actively maintained by a specific cellular transcriptional, signaling, and metabolic program. Despite the prevalence of nondividing quiescent cells in the body, quiescence is relatively understudied, and little is understood about the signaling mechanisms that govern it. Studies have found that upon entry into quiescence, the cell undergoes a reorganization of its cellular structure. The actin cytoskeleton is relocated into actin bodies [12]; yeast relocates its proteasome from the nucleus into storage chambers in the cytoplasm [13],, and many cytoplasmic proteins are compartmentalized into molecular complexes [14]. There are several in vitro culture systems of quiescence that use (combinations of, or, in isolation) contact inhibition, mitogen deprivation, nutrient starvation, or loss of adhesion [3–5, 15, 16]. Though each approach allows cell cycle exit, they induce different cellular programs such as mechanotransduction regulatory mechanisms after contact inhibition or mitogen-dependent reduction in proliferation signaling cascades during mitogen removal [3–5, 8, 16]. Interestingly, cell model systems that relied on varying the duration of contact inhibition resulted in the generation of heterogenous populations of both deep and shallow quiescent cells, with increased duration being correlated with increased depth of quiescence [4, 5, 8, 16–20]. Similarly, the duration of reduced mitogen correlated with the depth of the quiescent program [5, 8, 16]. Another source of heterogeneity is the variability in cell samples. As regular cell lines are usually of tumor origin, they have lost contact inhibition and/or sensitivity to mitogen removal and do not enter G0 [21]. Primary cells are commonly used instead, but as they enter replicative senescence after few passages in vitro [22], new cell samples are required regularly. The genetic diversity of cell donors typically leads to heterogeneity between experiments. These limitations can be overcome by the use of cells immortalized by the human telomerase reverse transcriptase (hTERT). Such cell lines retain primary characteristics for several hundreds of passages in vitro, allowing for the reproducibility of experiments over long periods of time [23, 24]. There are now

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hTERT-immortalized cells from 16 human tissues available commercially. Accordingly, we have generated a quiescent cell culture model that uses hTERT-immortalized cell lines in combination to both long-duration contact inhibition and an extended period of mitogen removal. This approach has yielded a reproducible and largely homogenous cell culture system with characteristics of long-term quiescence. Here, we employ this culture model to measure endocytosis during deep quiescence. Endocytosis is the process by which cells acquire substances from outside the cell and internalize surface membrane proteins [25]. Endocytosis is required by all eukaryotic cells for communication with their environment, internalization of micronutrients, and turnover of cell surface components. It not only supports the steady-state distribution of cell surface receptors but also regulates their activity by mediating their removal from the cell surface and degradation by lysosomes. As such, endocytosis plays key roles in biological processes such as synaptic transmission and signal transduction and in controlling developmental processes regulating cell fate [26–28]. The process occurs through invaginations of the plasma membrane forming endocytic vesicles that carry the “cargo” molecules (nutrients or receptors). The detachment of endocytic carriers relies on the GTPase Dynamin or other scission mechanisms [29]. Furthermore, endocytosis is also exploited by many pathogens (toxins, viruses, bacteria) as portal of entry. Mis-regulation of endocytosis through mutation or other means can cause a broad range of diseases including cancer, atherosclerosis, neurodegeneration, and lysosomal storage diseases [26, 30, 31]. There are several parallel endocytic pathways. The most thoroughly described of them is clathrin-mediated endocytosis (CME), which is the dominant uptake mechanism supporting cellular homeostasis [32, 33]. This constitutive process has a molecular hallmark attributed to the dependency on clathrin, a triskelionshaped scaffold protein comprising three light and three heavy chains [32, 33]. Cells also feature several clathrin-independent pathways of endocytosis (CIE) that allow them to take up membrane and extracellular components in diverse geometries at both macro- and microscales [34, 35]. Generally, CIE are involved in cellular processes outside of a housekeeping role, including rapid removal of activated receptors for the plasma membrane, bulk lipid or protein internalization, and cell morphology events like migration, polarization, and cell spreading [34, 35]. CIE pathways can be further subdivided by the scales and morphology of the membrane cargo carriers that they internalize, including large-membrane (0.2 to >10 μm) macropinosomes, tubular carriers as occurs in activated pathways like fast-endophilin-mediated endocytosis (FEME), and small (50–200 nm) micropinosomes such as CLIC/GEEC

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[34]. Each of these separate CIE pathways are further segregated by their dependency on dynamin and the cargoes they transport [34– 36]. Given the fundamental nature of endocytosis and the breath of the biological processes that it mediates, it is expected that its activity varies along the cell cycle, including quiescence. However, evidence to date is biased toward permanently proliferating cells as the vast majority of studies used cancer cell lines that cannot enter quiescence [21]. Here, we present a high-throughput imaging approach, coupled to open-source automated image analysis, that allows for the analysis of various endocytic pathways in both actively cycling cell populations and in long-term quiescent populations. The approach ensures that each stage of the cell cycle is represented with sufficient cell numbers (many thousands) to allow for robust statistical analysis. Lastly, the cellular models and analysis approaches presented are readily suitable to be adapted to other cellular events beyond endocytosis that may vary across the full spectrum of the proliferative cell cycle.

2

Materials 1. Primary cells (e.g., primary fibroblasts, HUVEC) or hTERTimmortalized normal cells (e.g., RPE1) (see Note 1). 2. Serum-containing culture medium appropriated for cells used in Subheading 2, Item 1 (see Note 2). 3. Serum-free culture medium appropriated for cells used in Subheading 2, Item 1 (see Note 3). 4. Enzyme-free cell detachment solution (see Note 4). 5. Cell counter. 6. Glass- or cycloolefin-bottom 96-well microplates (see Note 5). 7. Phosphate-buffered saline (PBS) buffer: 137 mM NaCl, 2.7 mM KCl, 12.5 mM Na2HPO4, 2 mM KH2PO4 equilibrated at pH 7.4. 8. PBS++: containing 0.9 mM CaCl2 and 0.5 mM MgCl2 (see Note 6). 9. Treatments being investigated (inhibitor drugs, siRNA, plasmids, etc.). Here, we provide guidance for the use of chemical inhibitors blocking CME (Pitstop 2, stock 50 mM in DMSO; see Note 7), Dynamin-dependent endocytosis (Dyngo-4a, stock 30 mM in DMSO; chlorpromazine stock 20 mM in water (see Note 8); Dynole 34-2, stock 10 mM in DMSO; Indole 24, stock 30 mM in DMSO; Quinone 45, stock 30 mM in DMSO; Pyrimidyn 7, stock 30 mM in DMSO; MitMAB, stock 10 mM in water; Aminopyrimidine, stock 20 mM in DMSO, or Dynasore, stock 50 mM in DMSO),

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and macropinocytosis (EIPA, stock 50 mM in DMSO; Rottlerin, stock 5 mM in DMSO), as well as RNA interference of AP2 (siRNA oligonucleotides (CCGCCAGAUGGAGAGUUUGAGCUUA and UAAGCUCAAACUCUCCAUCUGGCGG) reconstituted in milliQ water to a 20 μM stock (see Note 9). 10. Transfection reagent as appropriate for the cell type used. 11. Low-serum transfection medium, used to form RNAiLipofectamine complexes. 12. Fluorescently labeled ligands to monitor CME (e.g., AlexaFluor488-, AlexaFluor555-, or AlexaFluor647-labeled transferrin; DiI- or DiO-labeled low-density lipoprotein (LDL)), Dynamin-dependent endocytosis (e.g., AlexaFluor488-, AlexaFluor555-, or AlexaFluor647-labeled epidermal growth factor (EGF)), and macropinocytosis (e.g., AlexaFluor555-, AlexaFluor647-, or TRITC-labeled Dextran of various sizes (3,000 to 70,000 kDa), Lucifer yellow, DQ Green BSA) (see Note 10). 13. Antibodies suitable for endocytic feeding assays (e.g., anti-TfR or anti-EGFR antibody) (see Note 11). 14. Ligand uptake assay (LUA) medium: regular serum-free or full-growth medium or α-MEM without phenol red supplemented with 20 mM Hepes, pH 7.2, and 1% BSA. 15. Stripping buffer 1 (pH 5.5): 150 mM NaCl, 100 mM glycine, 5 mM KCl, 1 mM CaCl2, 1 mM MgCl2 adjusted to pH 5.5. 16. Stripping buffer 2 (pH 2): 150 mM NaCl, 0.2 M acetic acid, 5 mM KCl, 1 mM CaCl2, 1 mM MgCl2, adjusted to pH 2. 17. Paraformaldehyde (PFA), diluted to 4% in PBS. 18. PBS containing 50 mM NH4Cl. 19. Hoechst 33342 (stock 10 mg/mL in water) and DAPI (stock 1 mg/mL in water) (see Note 12). 20. Fluorescent widefield high-content screening microscope equipped with 20
or 40
air objective (numerical aperture ¼ 0.45) and an environmental temperature and CO2 control module (see Note 13). 21. Saponin. 22. Microscopy slide mounting solution (Mowiol or equivalent), containing antifading agent (DABCO or equivalent).

3

Methods 1. Grow cells in full medium at 37  C. 2. Passage cells when they reach ~70% confluency (see Note 14).

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3.1 Maintenance of Exponentially Growing Cells

(a) Detach cells using nonenzymatic cell detachment solution (10 min, 37  C) (see Note 4). (b) Seed cells at 0.75 to 1.5
104 cells/cm2 (or 1:10 to 1:5 volume dilution ratio) (see Note 15). 3. Change medium every 2 days (see Note 16). 4. Two days before experiments, growing cells were seeded in the appropriate dish or plate format at a density of 1.5
104 cells/ cm2.

3.2 Induction of Cellular Quiescence

1. Use exponentially growing cell cultures at ~70% confluency (see Note 14). 2. Detach cells using enzyme-free cell detachment solution (10 min, 37  C) (see Note 4). 3. Count cells and seed them in the appropriate dish or plate format at a density of 1.4
104 cells/cm2 (see Note 17). 4. Change medium (full serum) every 2 days (see Note 18). 5. Grow cells in full medium for at least 7 days until they reach confluence and form a homogeneous monolayer (see Note 19). 6. Change medium to growth factor-free (serum-free) medium (see Note 20). 7. Change medium (serum-free) every 2 days. 8. Maintain cells for at least 10 days (thus, at least 17 days from seeding) to induce deep quiescence (Fig. 1) (see Note 21).

3.3 Inhibiting Endocytosis During Cellular Quiescence

3.3.1 CME Inhibition by RNA Interference Growing Cells

Endocytosis can be inhibited by gene ablation, RNA interference (RNAi), overexpression of dominant-negative proteins, protein relocation, or addition of small compounds inhibitors. Because of the many entry routes into cells, endocytic pathways cannot be all blocked at once—a perturbation that would be highly toxic to most, if not all, cells. To date, only CME, macropinocytosis, CLIC/GEEC, FEME, and Dynamin-dependent endocytosis (which includes CME and other pathways such as FEME) can be inhibited with some specificity. Owing to the difficulty of transfecting and expressing DNA plasmids in quiescent cells, we chose to present the inhibition of CME using AP2 RNAi and Pitstop 2 and blockage of macropinocytosis and Dynamin-mediated endocytosis using small compound inhibitors. 1. Six hours prior to transfection, seed growing cells at 6.9
103 cells/cm2 in 96-well plates. 2. For each well, dilute 0.17 μL of Lipofectamine RNAiMAX transfection reagent in 4.83 μL OptiMEM and 0.0675 μL of μ2-adaptin siRNA in 4.93 μL OptiMEM, and incubate 5 min at room temperature.

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Fig. 1 Induction of long-term quiescence; bar, 50 μm

3. Mix diluted RNAiMAX with diluted siRNA, and incubate for 20 min at room temperature. 4. Add 10 μL transfection mix to each well containing cells in 100 μL growth medium. 5. Repeat the transfection 24 h after step 4. 6. Change the medium after 48 h. 7. Use the cells 72 h after step 4. Quiescent Cells

1. Five days prior to transfection, seed growing cells at 1.4
104 cells/cm2 in 96-well plates (see Note 22). 2. For each well, dilute 0.85 μL of Lipofectamine RNAiMAX Transfection Reagent in 4.15 μL OptiMEM and 0.1 μL of μ2-adaptin siRNA in 4.9 μL OptiMEM, and incubate 5 min at room temperature. 3. Mix diluted RNAiMAX with diluted siRNA, and incubate for 20 min at room temperature.

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4. Add 10 μL transfection mix to each well containing 100 μL growth medium. 5. Change the medium (full serum) after 24 h. 6. Repeat the transfection 48 h after step 4. 7. Change the medium to serum-free 10 days after step 1. 8. Change the medium (serum-free) every 48 h. 9. Use the cells ~12 days after step 4. 3.3.2 CME Inhibition by Pitstop 2

Using the cell-seeding procedure detailed in Subheadings 3.1 and 3.2, treat growing and quiescent cell cultures as follows: 1. Dilute Pitstop 2 to 30 μM into the respective culture media. 2. Replace the culture media with mix from step 1, and incubate at 37  C for the appropriate duration (a minimum of 10 min and maximum of 4 h is recommended) prior to the endocytic assays.

3.3.3 Dynamin Inhibition by Small Inhibitors

Dynamin is inhibited in cells by several small compound inhibitors at the following indicative working concentrations: Dyngo-4a 4–10 μM [37], chlorpromazine 18 μM [38], Dynole 34-2 10 μM [39] (see Note 23), Indole 24 2 to 5 μM [40], Quinone 45 50 μM [41], Pyrimidyn 7 10 to 30 μM [42], MitMAB 30 μM [43], Aminopyrimidine 10 to 50 μM [44], or Dynasore 80 μM [45] (see Note 24). It is good practice to use at least two independent inhibitors to limit off-target effects (see Note 25). Inhibiting Dynamin has a broad effect on endocytosis, consistent with its function in several endocytic pathways (including CME and FEME). Using the cell-seeding procedure detailed in Subheadings 3.1 and 3.2, treat growing and quiescent cell cultures as follows: 1. Dilute the inhibitors to the appropriate concentration into serum-free medium (for both growing and quiescent cells) (see Note 26). 2. Wash cells in serum-free medium. 3. Replace the culture media with mixes from step 1, and incubate at 37  C for the appropriate duration (a minimum of 10 min and maximum of 4 h is recommended) prior to the endocytic assays.

3.3.4 Macropinocytosis Inhibition by Small Inhibitors

Micropinocytosis (fluid-phase uptake) and macropinocytosis are inhibited by Na+/H+ exchange inhibitor ethyl-isopropyl amiloride (EIPA) at 10–25 μM [46, 47] or by Rottlerin at 2–10 μM [48]. Using the cell-seeding procedure detailed in Subheadings 3.1 and 3.2, treat growing and quiescent cell cultures as follows: 1. Dilute the inhibitors to the appropriate concentration into the respective culture media.

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2. Replace the culture media with mixes from step 1, and incubate at 37  C for the appropriate duration (a minimum of 10 min and maximum of 4 h is recommended) prior to the endocytic assays. 3.4 Endocytic Assays in Quiescent Cells

Endocytosis of receptors can be measured either by the intracellular entry of antibody recognizing their ectodomains (antibody feeding assay) or by the uptake of their ligands. For CME, we recommend fluorescently labeled transferrin (50–200 μg/mL) and LDL (5–15 μg/mL) or anti-TfR antibody suitable for feeding assay (0.1–10 μg/mL); for Dynamin-mediated endocytosis, fluorescently labeled EGF (50 ng/mL) or anti-EGFR antibody suitable for feeding assay (0.1–10 μg/mL, together with 50–100 ng/mL unlabeled EGF); for micropinocytosis, Lucifer yellow (5 mg/mL); and for macropinocytosis, fluorescently labeled Dextrans (0.5–10 mg/mL depending on size and fixation) and fluorescently labeled BSA or DQ-BSA (100 μg/mL) (see Note 27).

3.4.1 Uptake of Fluorescently Labeled Fixable Ligands

For fluorescently labeled Tf, LDL, and EGF that can be fixed by 4% PFA, do as follows: Cells seeded on glass-bottom microplates should be prepared according to Subheadings 3.1, 3.2, and 3.3, as appropriate. 1. Pre-warm (37  C for 1 h) ligand uptake assay (LUA) medium (see Note 28). 2. Dilute fluorescently labeled ligand to the desired concentration in warm LUA medium (see Note 29). 3. Take cells out of the incubator, swiftly aspirate culture medium, and replace it with pre-warmed LUA medium containing ligand, return the cells to the 37  C incubator, and start a timer (see Note 30). 4. After desired time has elapsed, remove the samples from the incubator, and place them on a water-ice tray (see Note 31). 5. Aspirate LUA medium, and wash the samples two times with ice-cold PBS++. 6. Wash the samples two times (2 min each) with Stripping Buffer 1 (see Note 32). 7. Wash the cells three times with ice-cold PBS++ (see Note 33). 8. Add ice-cold fixative solution (4% PFA in PBS), and incubate on ice for 5 min and another 15 min at room temperature (see Note 34). 9. Wash the fixed cells three times with PBS containing 50 mM NH4Cl (see Note 35). 10. Wash the samples three times with PBS, and immunostain or store at 4  C as required (see Subheading 3.5).

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3.4.2 Uptake of Fluorescently Labeled Non-fixable Ligands

For fluorescently labeled Dextrans, Lucifer Yellow, and DQ-BSA that cannot be fixed, do as follows: Cells seeded on glass- or cycloolefin-bottom microplates should be prepared according to Subheadings 3.1, 3.2, and 3.3 as apropriate. 1. Pre-warm (37  C for 1 h) ligand uptake assay medium (see Note 28). 2. Dilute fluorescently labeled ligand to the desired concentration in warm ligand uptake assay (LUA) medium (see Note 29). 3. Take cells out of the incubator, swiftly aspirate culture medium, and replace it with pre-warmed LUA medium containing ligand, return the samples to the 37  C incubator, and start a timer (see Note 30). 4. 20 min before the end of the incubation time, add Hoechst 33342 (final concentration 2.5 μg/mL), and return samples to 37  C (see Note 36). 5. After desired time has elapsed, remove the samples from the incubator and place them on a water-ice tray, aspirate assay medium, and wash the samples five times with ice-cold PBS++ (see Note 31). 6. Exchange PBS to Imaging Medium, and image the cells imediately (see Note 37).

3.4.3 Antibody Feeding Assays

Cells seeded on glass- or cycloolefin-bottom microplates should be prepared according to Subheadings 3.1, 3.2, and 3.3, as appropriate. 1. Pre-warm (37  C for 1 h) antibody uptake assay medium (see Note 28). 2. Dilute antibodies to the desired concentration (e.g., 5 μg/mL for an antibody against the ectodomain of EGFR) in LUA medium (see Note 29). 3. Take cells out of the incubator, swiftly aspirate culture medium, and replace it with pre-warmed assay medium containing antibodies, return the cells to the 37  C incubator, and start a timer (see Note 30). 4. After the desired time has elapsed, remove the samples from the incubator, and place them on a water-ice tray. 5. Aspirate LUA medium, and wash the samples three times with ice-cold PBS++ (see Note 31). 6. Wash the cells three times with Stripping Buffer 2 for 2 min (see Note 38). 7. Wash the cells three times with ice-cold PBS++ (see Note 33).

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8. Add ice-cold fixative solution (4% PFA in PBS), and incubate on ice for 5 min and another 15 min at room temperature (see Note 34). 9. Wash the fixed cells three times with PBS containing 50 mM NH4Cl (see Note 35). 10. Wash the samples three times with PBS, and immunostain or store at 4  C as required (see Subheading 3.5). 3.4.4 Cell Surface Receptor Labeling

Correcting ligand/antibody uptake measurements with cell surface receptor availability is required to normalize for receptor abundance between growing and quiescent cells (a ligand might be less internalized during G0 because the cell surface levels of its receptor are decreased, instead of because the endocytic pathway is downregulated). Cells seeded on glass- or cycloolefin-bottom microplates should be prepared according to Subheadings 3.1, 3.2, and 3.3, as appropriate. 1. Remove the samples from the incubator, and place them on a water-ice tray (see Note 39). 2. Wash the cells three times with ice-cold PBS++ (see Note 39). 3. Keep cells on the water-ice tray, and incubate them with Imaging Medium containing 1% BSA and the primary antobdies at the desired concentrations. 4. Incubate the samples for 90 min on a water-ice tray in a cold room (see Note 39). 5. Wash the cells three times with ice-cold PBS++ (see Note 39). 6. Add ice-cold fixative solution (4% PFA in PBS), and incubate on ice for 5 min and another 15 min at room temperature (see Note 34). 7. Wash the fixed cells three times with PBS containing 50 mM NH4Cl (see Note 35). 8. Wash the samples three times with PBS, and immunostain or store at 4  C as required (see Subheading 3.5).

3.5 Fixation and Immunofluorescence Labeling

1. Wash the samples three times with PBS. 2. Incubate the samples with blocking buffer containing 5% BSA and 0.1% Saponin for 30 min at room temperature (see Note 40). 3. Remove blocking buffer, and incubate the cells with the relevant primary antibodies diluted in PBS containing 5% BSA and 0.1% Saponin for 90 min at room temperature or overnight at 4  C. 4. Wash samples three times with PBS (5 min incubations) containing 0.1% Saponin.

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5. Incubate cells with the relevant secondary antibodies and DAPI (2 μg/mL) diluted in PBS containing 5% BSA and 0.1% Saponin for 45 min in the dark, at room temperature (see Note 41). 6. Wash samples three times (5 min incubations) with PBS containing 0.1% Saponin. 7. Wash samples twice with PBS and twice with milliQ water. 8. Exchange the water for antifading mounting solution (containing Mowiol and DABCO or equivalent, 50–100 μL per well). 9. Image immediately, or incubate the samples overnight at room temperature before long-term storage at 4  C. 3.6 Image Acquisition by High-Throughput Automated Widefield Microscopy

The protocol below is for image acquisition of glass or cycloolefinbottom 96-well plates on an ImageXpress Micro XL Widefield High-Content Screening System (Molecular Devices) or equivalent. The protocol can be adapted for manual image acquisition, regular microscopy slides, and confocal or super-resolutionmicroscopy, as appropriate (Fig. 2). 1. Prior to imaging, calibrate plates to allow for 96-well plate and bottom autofocus setup (Fig. 3, step 1).

Fig. 2 Adaptations for analysis of high-throughput experiments in 96-well plates

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Fig. 3 Plate setup and image acquisition

2. Select objective (see Note 42) (Fig. 3, step 2). 3. Select the number of locations and sites to be imaged (Fig. 3, step 3). 4. Set up the acquisition loop for each image (Fig. 3, step 4): (a) Select laser-based focusing on plate and well bottom for each image. (b) Select wavelength channels for excitation and emission appropriate for your fluorophore dyes. (c) Select the channel serving as focus offset for each other channel. (d) Determine the image-based focus and exposure times for each channel.

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Fig. 4 Regular expression to extract metadata information from image file names in CellProfiler

5. Apply a focus journal after each image acquisition to account for unevenness in plate design and focus planes (Fig. 3, step 5) (see Note 43). 6. Run the image acquisition for the wells selected. 3.7 Automated Image Analysis

We recommend analyzing the images acquired by confocal or highthroughput microscopy with the open-source software CellProfiler 2.2.0 [49, 50]. Alternative analysis software or routines can be used instead. Import image sets (one image set consists of up to four separate images, one for each wavelength) into CellProfiler together with metadata information about experiment name (Fig. 4), well and site number, and wavelength channel extracted with regular expressions.

3.7.1 Cell Segmentation and Endosome Identification

The workflow of a CellProfiler pipeline-segmenting cells and endosomes is shown in Fig. 5. 1. Treat the grayscale images for each wavelength channel separately. 2. Calculate an illumination function and background-subtract it from the raw images to account for uneven illumination during image acquisition (Fig. 5, step 1). 3. Rescale illumination-corrected images (on a scale of 0–1) so that faint signals are visible for object identification (Fig. 5, step 2). 4. On rescaled images, identify nuclei labeled with DAPI or Hoechst 33342 as primary objects (Fig. 5, step 3). 5. Identify cell bodies labeled with a cytoplasmic marker (such as GAPDH or F-actin) as secondary objects emanating from a primary object (Fig. 5, step 4). 6. Create a mask on the identified cell bodies to eliminate areas not occupied by cells from further analysis (Fig. 5, step 5). 7. Within the mask, enhance speckles (which are small circular areas of increased intensity compared to their immediate environment) to distinguish them from background haze. Additionally, this step eliminates large areas of signal caused by antibody aggregation or surface-bound antibody that was not efficiently stripped (Fig. 5, step 6).

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Fig. 5 CellProfiler workflow to segment cells and identify internalized ligand or receptor puncta

8. Use the speckle-enhanced images to identify endosomes as puncta of a defined pixel size and minimum intensity (usually 4–15 pixels diameter, dependent on stringency) (Fig. 5, step 7). 9. Measure mean intensity and total area of puncta identified in illumination-corrected images created in step 2 (Fig. 5, step 8).

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Fig. 6 Object-based colocalization using CellProfiler

10. For quantitation, subtract mean fluorescence of puncta per image by the mean fluorescence of cells per image in blank images (cells not subjected to ligand uptake but else treated in the same way). Finally, multiply by the total area of puncta per cells to get the sum of the ligand internalized into endosomes. 3.7.2 Object-Based Colocalization Analysis

The workflow of a CellProfiler pipeline identifying endosome colocalization is shown in Fig. 6. The pipeline is available on the CellProfiler website (https://cellprofiler.org/published-pipelines). This analysis is only suitable for images acquired by confocal microscopy. 1. Correct (illumination correction) and rescale images, enhance speckles, and identify endosomes as in Subheading 3.7.1. 2. Shrink identified puncta either to a central pixel or by a specified number of pixels, according to the desired analysis stringency (Fig. 6, step 3).

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3. Identify the colocalization of shrunk puncta in different channels (images) by relating overlapping objects in a child-parent relationship and filter children with a parent or vice versa (Fig. 6, step 4). 4. To relate the intensities measured to the original-size endosomes, relate the shrunken puncta with their parent (or child) back to the puncta identified initially (Fig. 6, step 5). 5. Measure the percentage of colocalization by relating the total intensity of colocalized signals and total amount of identified puncta in illumination-corrected images. 3.8 Statistical Analysis

An advantage of high-content imaging is the measurement of high numbers (multiple thousands) of cells, giving robust statistics. 1. Test for Gaussian distribution of the data using D’Agostino and Pearson omnibus normality test or equivalent. 2. For samples that follow Gaussian distribution: Test for statistical significance using Student’s or Welch’s unpaired two-tailed t-test (two sample groups), one-way ANOVA and Dunnett’s test for multiple comparisons (more than two sample groups), or two-way ANOVA with Tukey’s test for multiple comparisons (more than two sample groups with multiple parameters per group), as appropriate. 3. For samples that do not follow Gaussian distribution: Test for statistical significance using Mann-Whitney U-test (two-sample groups), Kruskal-Wallis test with Dunn’s multiple comparison (more than two sample groups), or Friedman test with Dunn’s test for multiple comparisons (more than two sample groups with multiple parameters per group), as appropriate.

3.9 Representative Result

4

Figure shows EGF uptake (50 ng/mL AlexaFluor647-EGF for 15 min at 37  C) into growing and quiescent cells. Approximately 400,000 and 60,000 intracellular EGF puncta were identified and quantified in 10,000 and 19,000 growing and quiescent cells, respectively, corresponding to an average of 40 puncta per growing and 3 puncta per quiescent cell (Fig. 7).

Notes 1. A wide variety of cells may be used as long as they maintain cell cycle exit upon contact inhibition (it is not the case for commonly used tumor-derived cell lines). The methods described here use human normal, diploid hTERT-RPE1 cells but are easily adapted for other primary or hTERT-immortalized cell types.

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Fig. 7 Quantification of 15 min EGF uptake in growing and quiescent cells. Scale bar: 50 μm

2. Growing cells are cultured in full serum-containing medium (e.g., 10% FBS or equivalent). 3. Quiescent cells survive for long periods of time (months) in growth factor-free medium (i.e., serum-free) containing regular levels of amino acids, glucose, and other supplements. 4. Detaching cells with trypsin-based solutions is possible although less desirable as it shaves cells from a significant proportion of their cell surface receptors. 5. The protocol can be adapted to any other size and type of tissue culture containers. 6. Presence of Ca2+ and Mg2+ is required for cell adhesion and survival until fixation. 7. Other chemical inhibitions of CME such as hypertonic shock (0.45 M sucrose), cytosol acidification [51], potassium depletion, monodansylcadaverine, or phenylarsine oxide [52–56], even though widely used in the literature, are not recommended as they affect clathrin-independent endocytosis, including FEME, as well [57]. 8. Chlorpromazine was initially believed to be a clathrin and/or AP2 inhibitor [56] but was recently found to inhibit Dynamin instead [38]. 9. RNA interference of AP2 is preferable to that of clathrin as the latter has many functions beside endocytosis [58]. Gene editing using CRISPR/Cas9 is also possible. 10. The choice of fluorophore depends on the sensitivity of the microscope used. We found that Alexa647-labeled ligands have the highest signal-to-noise ratio. 11. Antibodies suitable for feeding assays must bind to the extracellular parts of the receptor of interest.

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12. While both intercalating dyes stain DNA in a stoichiometric manner, Hoechst 33342 is cell-permable (and thus can be used on live cells), whereas DAPI is not. 13. The protocol can be easily adapted to any kind of widefield, confocal, or super-resolution microscopes, as appropriate. 14. We found that passaging primary and hTERT-immortalized cells before they reach such confluency ensures exponential growth while minimizing subpopulations of quiescent cells (happening when patches of cells become confluent). 15. We typically passage growing cells every 2 days (highest seeding density provided). Seeding cells at the lowest density will allow for ~3 days until the next passage. 16. Growing cells need full-serum medium. 17. We found that seeding cells at this density ensures ordinate formation of homogeneous monolayers. 18. Cells must be grown in presence of growth factor-containing (full-serum) medium until tight monolayers are formed. 19. The formation of tight and well-organized monolayers (i.e., where cells are oriented in the same direction) is important to induce deep quiescence. 20. Once monolayers are formed, cells can be maintained in growth factor-free (serum-free) medium. Primary cells will enter apoptosis if growth factors are removed prior to cell confluency. 21. We found that 10 days growth factor removal after monolayer establishment is enough to induce deep quiescence. 22. We found that siRNA oligos transfection and RNA interference is more efficient while the cells are entering quiescence. Once in G0, protein levels of RNAi targets remain depressed for long periods (multiple days). 23. Dynole 32-1 is a negative control compound for Dynole 34-1 [39]. 24. Although not as potent or specific as the ones selected, other Dynamin inhibitors such as OctMAB, Pro-Myristic Acid, Rhodadyn C10, or Iminodyn-22 [39, 43, 59] can be used as well. 25. It is best practice to use small compound inhibitors against different domains of Dynamin (e.g., Dyngo-4a or Dynole 34-2 targeting the G-domain and MitMAB, OctAB, or Pro-Myristic Acid targeting the PH domain). 26. Many small compound inhibitors are hydrophobic and quenched by serum (mostly albumin present therein), thereby reducing their active concentration.

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27. Lucifer yellow is a fluid-phase marker. Dextrans of various sizes (from 3,000 to 2,000,000+ kDa) can be used to discriminate between small micropinosomes and large macropinosomes. DQ-BSA only fluoresces when it reaches degradative endolysosomes, thereby controlling for uptake of the probe. 28. Pre-warming ligand/antibody uptake medium is required to maintain the temperature of the cells as close to 37  C as possible throughout the assay. 29. Small compound inhibitors must be present with the ligands for the relevant samples. 30. Incubation times can vary from 1 min to 60 min, as required. To measure endocytic rates (expressed as amount of ligand per unit of time), several lengths of incubations must be measured. 31. The use of cold temperatures at the end of the assay is appropriate as the cells will be fixed shortly after. Adding ice-cold PBS++ stops any trafficking within seconds. Using ice + water mix to cool the plates/dishes during the subsequent steps ensures a better temperature exchange and cooling than using ice only. 32. Washes with a mildly acidic stripping buffer (“Stripping Buffer 1,” pH 5.5) remove cell surface ligands that were not internalized after incubation (internalized ligands are protected within endocytic carriers or endosomes). 33. Washes with PBS are required for raising the pH back to ~7.4 after washes with stripping buffers. 34. Cold fixation is required to avoid that endocytosis resumes unwittingly. 35. Washes with 50 mM NH4Cl in PBS are required to inactivate any residual reactive PFA. 36. Staining DNA with cell-permeable Hoechst 33342 is important for automated cell identification (Subheading 3.7). 37. Imaging should be done at 37  C if image acquisition can be done within 10 min. Do the imaging at 4  C otherwise (to slow down endosomal recycling and degradation of the ligands). 38. Washes with an acidic stripping buffer (“Stripping Buffer 2,” pH 2) remove cell surface antibodies that were not internalized after incubation. Note that cell surface stripping of, particularly high-affinity, antibodies requires lower pH than that of endocytic ligands. 39. Low temperature is required for cell surface staining to stop receptor trafficking. 40. We typically use Saponin to permeabilize cells, but other means (Triton X-100, cold methanol) can be used as well.

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41. Fluorescently labeled secondary antibodies toward the species of internalized antibodies must be included in the immunostaining procedure (e.g., AlexaFluor-488-coupled goat antirabbit secondary antibodies to label rabbit anti-EGFR antibodies internalized during the feeding assay). Counterstaining of protein(s) of interest, DNA (using DAPI or DRAQ5), and/or actin cytoskeleton (phalloidin) using other fluorophores than those coupled on the internalized ligand can be considered. 42. 20
objectives are typically sufficient to identify single endosomes while acquiring enough cells per field of view. 43. The journal encoded a z-stack (e.g., four images with 2 μm spacing, set manually) from which the image with the best focus is saved as final acquisition.

Acknowledgments C.H. was supported by a studentship from the British Heart Foundation (FS/14/20/30681). K.McG. was supported by the Biotechnology and Biological Sciences Research Council (BBSRC) and a UL-Health Research Institute Seed Award. E.B. was a Biotechnology and Biological Sciences Research Council (BBSRC) David Phillips Research Fellow and a Lister Institute Research Fellow. Part of this work was supported by a BBSRC grant (BB/R0155X/1) and a Birkbeck Wellcome Trust Institutional Strategic Support Fund (ISSF) grant to E.B. References 1. Daignan-Fornier B, Sagot I (2011) Proliferation/quiescence: the controversial “Allerretour”. Cell Div 6:10 2. Herman PK (2002) Stationary phase in yeast. Curr Opin Microbiol 5:602–607 3. Gos M, Miloszewska J, Swoboda P, Trembacz H, Skierski J, Janik P (2005) Cellular quiescence induced by contact inhibition or serum withdrawal in C3H10T1/2 cells. Cell Prolif 38:107–116 4. Coller HA, Sang L, Roberts JM (2006) A new description of cellular quiescence. PLoS Biol 4: e83 5. Lemons JMS, Feng X-J, Bennett BD, LegesseMiller A, Johnson EL, Raitman I, Pollina EA, Rabitz HA, Rabinowitz JD, Coller HA (2010) Quiescent fibroblasts exhibit high metabolic activity. PLoS Biol 8:e1000514 6. Fuge EK, Braun EL, Werner-Washburne M (1994) Protein synthesis in long-term

stationary-phase cultures of Saccharomyces cerevisiae. J Bacteriol 176:5802–5813 7. Gray JV, Petsko GA, Johnston GC, Ringe D, Singer RA, Werner-washburne M (2004) Sleeping beauty: quiescence in Saccharomyces cerevisiae. Microbiol Mol Biol Rev 68:187–206 8. Gookin S, Min M, Phadke H, Chung M, Moser J, Miller I, Carter D, Spencer SL (2017) A map of protein dynamics during cell-cycle progression and cell-cycle exit. PLoS Biol 15:e2003268 9. Valentin M, Yang E (2008) Autophagy is activated, but is not required for the G0 function of BCL-2 or BCL-xL. Cell Cycle 7:2762–2768 10. Crissman HA, Darzynkiewlcz Z, Tobey RA, Steinkamp JA (1985) Normal and perturbed Chinese hamster ovary cells: correlation of DNA, RNA, and protein content by flow cytometry. J Cell Biol 101:141–147

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24. Kogan I, Goldfinger N, Milyavsky M, Cohen M, Shats I, Dobler G, Klocker H, Wasylyk B, Voller M, Aalders T, Schalken JA, Oren M, Rotter V (2006) hTERTimmortalized prostate epithelial and stromalderived cells: an authentic in vitro model for differentiation and carcinogenesis. Cancer Res 66:3531–3540 25. Doherty GJ, McMahon HT (2009) Mechanisms of endocytosis. Annu Rev Biochem 78:857–902 26. Sigismund S, Confalonieri S, Ciliberto A, Polo S, Scita G, Di Fiore PP (2012) Endocytosis and signaling: cell logistics shape the eukaryotic cell plan. Physiol Rev 92:273–366 27. Sigismund S, Scita G (2018) The ‘endocytic matrix reloaded’ and its impact on the plasticity of migratory strategies. Curr Opin Cell Biol 54:9–17 28. Hinze C, Boucrot E (2018) Endocytosis in proliferating, quiescent and terminally differentiated cells. J Cell Sci 131:jcs216804 29. Renard HF, Johannes L, Morsomme P (2018) Increasing diversity of biological membrane fission mechanisms. Trends Cell Biol 28:274–286 30. McMahon HT, Boucrot E (2011) Molecular mechanism and physiological functions of clathrin-mediated endocytosis. Nat Rev Mol Cell Biol 12:517–533 31. Schmid SL (2017) Reciprocal regulation of signaling and endocytosis: implications for the evolving cancer cell. J Cell Biol 216:2623–2632 32. Kaksonen M, Roux A (2018) Mechanisms of clathrin-mediated endocytosis. Nat Rev Mol Cell Biol 19:313–326 33. Mettlen M, Chen P-H, Srinivasan S, Danuser G, Schmid SL (2018) Regulation of Clathrin-mediated endocytosis. Annu Rev Biochem 87:871–896 34. Ferreira APA, Boucrot E (2018) Mechanisms of carrier formation during Clathrinindependent endocytosis. Trends Cell Biol 28:188–200 35. Hemalatha A, Mayor S (2019) Recent advances in clathrin-independent endocytosis. F1000Res:8 36. Maldonado-Ba´ez L, Williamson C, Donaldson JG (2013) Clathrin-independent endocytosis: a cargo-centric view. Exp Cell Res 319:2759–2769 37. McCluskey A, Daniel JA, Hadzic G, Chau N, Clayton EL, Mariana A, Whiting A, Gorgani NN, Lloyd J, Quan A, Moshkanbaryans L, Krishnan S, Perera S, Chircop M, von Kleist L, Mcgeachie AB, Howes MT, Parton

Measuring Endocytosis During Proliferative Cell Quiescence RG, Campbell M, Sakoff JA, Wang X, Sun JY, Robertson MJ, Deane FM, Nguyen TH, Meunier FA, Cousin MA, Robinson PJ (2013) Building a better dynasore: the dyngo compounds potently inhibit dynamin and endocytosis. Traffic 14:1272–1289 38. Daniel JA, Chau N, Abdel-Hamid MK, Hu L, von Kleist L, Whiting A, Krishnan S, Maamary P, Joseph SR, Simpson F, Haucke V, McCluskey A, Robinson PJ (2015) Phenothiazine-derived antipsychotic drugs inhibit dynamin and Clathrin-mediated endocytosis. Traffic 16:635–654 39. Hill TA, Gordon CP, McGeachie AB, VennBrown B, Odell LR, Chau N, Quan A, Mariana A, Sakoff JA, Chircop M, Robinson PJ, McCluskey A (2009) Inhibition of dynamin mediated endocytosis by the Dynoles—synthesis and functional activity of a family of indoles. J Med Chem 52:3762–3773 40. Gordon CP, Venn-Brown B, Robertson MJ, Young KA, Chau N, Mariana A, Whiting A, Chircop M, Robinson PJ, McCluskey A (2013) Development of second-generation indole-based dynamin GTPase inhibitors. J Med Chem 56:46–59 41. MacGregor KA, Abdel-Hamid MK, Odell LR, Chau N, Whiting A, Robinson PJ, McCluskey A (2014) Development of quinone analogues as dynamin GTPase inhibitors. Eur J Med Chem 85:191–206 42. McGeachie AB, Odell LR, Quan A, Daniel JA, Chau N, Hill TA, Gorgani NN, Keating DJ, Cousin MA, Van Dam EM, Mariana A, Whiting A, Perera S, Novelle A, Young KA, Deane FM, Gilbert J, Sakoff JA, Chircop M, McCluskey A, Robinson PJ (2013) Pyrimidyn compounds: dual-action small molecule pyrimidine-based dynamin inhibitors. ACS Chem Biol 8:1507–1518 43. Quan A, McGeachie AB, Keating DJ, Van Dam EM, Rusak J, Chau N, Malladi CS, Chen C, McCluskey A, Cousin MA, Robinson PJ (2007) Myristyl trimethyl ammonium bromide and octadecyl trimethyl ammonium bromide are surface-active small molecule dynamin inhibitors that block endocytosis mediated by dynamin I or dynamin II? Mol Pharmacol 72:1425–1439 44. Odell LR, Abdel-Hamid MK, Hill TA, Chau N, Young KA, Deane FM, Sakoff JA, Andersson S, Daniel JA, Robinson PJ, McCluskey A (2017) Pyrimidine-based inhibitors of dynamin I GTPase activity: competitive inhibition at the Pleckstrin homology domain. J Med Chem 60:349–361 45. Macia E, Ehrlich M, Massol R, Boucrot E, Brunner C, Kirchhausen T (2006) Dynasore,

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Chapter 3 Measurements of Compensatory Endocytosis by Antibody Internalization and Quantification of Endocytic Vesicle Distribution in Adrenal Chromaffin Cells Mara Ceridono, Sylvette Chasserot-Golaz, Nicolas Vitale, Ste´phane Gasman, and Ste´phane Ory Abstract Plasma membrane proteins are amenable to endocytosis assays since they are easily labeled by reagents applied in the extracellular medium. This has been widely exploited to study constitutive endocytosis or ligand-induced receptor endocytosis. Compensatory endocytosis is the mechanism by which components of secretory vesicles are retrieved after vesicle fusion with the plasma membrane in response to cell stimulation and a rise in intracellular calcium. Luminal membrane proteins from secretory vesicles are therefore transiently exposed at the plasma membrane. Here, we described an antibody-based method to monitor compensatory endocytosis in chromaffin cells and present an image-based analysis to quantify endocytic vesicles distribution. Key words Compensatory endocytosis, Large dense core vesicle, Neuroendocrine cells, Chromaffin cells, Internalization, Dopamine-β-hydroxylase

1

Introduction A long-standing question is how cell plasma membrane homeostasis is maintained especially in cells with high exchange rate between pools of intracellular vesicles and the plasma membrane. Neurons and neuroendocrine cells are heavily challenged to maintain cell-tocell communication by releasing neurotransmitters and hormones through regulated exocytosis. To release their content upon cell stimulation, synaptic vesicles and secretory granules fuse with the plasma membrane resulting in a massive supply of vesicular membrane. Efficient compensatory endocytosis has to take place to maintain plasma membrane homeostasis and restore vesicular pools [1, 2]. Several accurate and sensitive methods are widely used to study exocytosis including electrophysiological and electrochemical approaches that measure changes in electrical properties

Florence Niedergang, Nicolas Vitale and Ste´phane Gasman (eds.), Exocytosis and Endocytosis: Methods and Protocols, Methods in Molecular Biology, vol. 2233, https://doi.org/10.1007/978-1-0716-1044-2_3, © Springer Science+Business Media, LLC, part of Springer Nature 2021

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of the plasma membrane and detect the released molecules, respectively [3–5]. Live cell imaging with the use of pH-sensitive fluorescent proteins which shine when pH increases upon fusion pore opening represents an alternative approach [6, 7]. Since plasma membrane recovers basal properties and fluorescent reporters are quenched upon acidification in endocytic compartments, such methods are, however, less suitable to follow the fate of proteins once the endocytic vesicle has formed. Therefore, the main challenge to study compensatory endocytosis is to specifically identify and track proteins that had been transiently exposed to the plasma membrane. Biochemical methods can give an overview of protein fate but often require large amounts of proteins that are not necessarily easily obtained, especially in model system with limited amount of cells such as mouse chromaffin cells, for example. Adrenal medulla chromaffin cells have been an instrumental model to study exocytosis and compensatory endocytosis. In contrast to synaptic vesicles which size range is below the resolution limits of conventional light microscopy, secretory granules of chromaffin cells allow for single-vesicle exocytosis and endocytosis analysis with conventional microscopy. By the use of specific antibodies that recognize luminal domain of secretory granule proteins, exocytic sites can be stained. Since vesicular trafficking and especially endocytosis require energy to proceed, chilling cells at 4
after exocytic stimulation temporarily blocks endocytosis, providing time to specifically label proteins of interest with antibodies. By replacing cells at physiological temperature, compensatory endocytosis gets started and antibodies bound to vesicular proteins are uptaken. We are providing here protocols for time course experiments and immunofluorescence analysis of endocytic vesicle containing antibodies directed against the luminal domain of the dopamine-β-hydroxylase (DBH), a secretory granule resident enzyme converting dopamine into noradrenaline. These protocols give access to spatiotemporal measurement of compensatory endocytosis in bovine chromaffin cells.

2 2.1

Materials Equipment

1. 12 mm coverslips cleaned and sterilized (see Note 1). 2. Tissue culture well plates with a diameter able to accept coverslips (typically 24 well plates or 4 well plates with diameter of 15 mm). 3. Ice bucket to transfer plates or maintain coverslips in plates at 4
C to stop vesicular trafficking during antibodies incubation. 4. Hot plate is used to maintain cells at 37
C during cell stimulation and endocytosis time course experiments.

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5. Forceps to transfer coverslips between plates. 6. Confocal microscope. 7. Open-source image analysis software (see Note 2). 8. Primary chromaffin cells cultured on coverslips (see Note 3). 2.2 Solution and Reagents

1. Locke’s solution: 140 mM NaCl, 4.7 mM KCl, 2.5 mM CaCl2, 1.2 mM MgSO4, 1.2 mM KH2PO4, 10 mM EDTA, 11 mM glucose, 0.57 mM ascorbic acid, and 15 mM HEPES, pH ¼ 7.5. 2. Nicotine solution: 1 M nicotine stock is prepared in Locke’s solution and conserved on ice protected from light. 10 μM nicotine in Locke’s solution is freshly prepared and warmed up to 37
C just before use. 3. High-potassium Locke’s solution: 85.7 mM NaCl, 59 mM KCl, 2.5 mM CaCl2, 1.2 mM MgSO4, 1.2 mM KH2PO4, 10 mM EDTA, 11 mM glucose, 0.57 mM ascorbic acid, and 15 mM HEPES, pH ¼ 7.5. 4. Phosphate buffer saline: 1 mM KH2PO4, 155 mM NaCl, 3 mM KH2PO4, pH 7.4. 5. Fibronectin from bovine plasma for coverslip coating: cleaned coverslips are incubated at least 2 h at 37
C in 50 μg/ml fibronectin diluted in water. 6. Cell culture medium: Bovine chromaffin cells are maintained in DMEM supplemented with 10% fetal bovine serum previously heat-inactivated for 30 min at 56
C, 1% glutamine, 1% cytosine arabinoside, and 0.1% fluorodeoxyuridine solutions to prevent fibroblasts proliferation. 7. Bovine serum albumin (BSA): 0.2% (w/v) BSA diluted in Locke’s solution (see Note 4). 8. Anti-dopamine-β-hydroxylase (DBH) antibodies (see Note 5). 9. Paraformaldehyde (PFA) solution: 16% PFA stock solution is diluted at 4% in PBS to fix cells. 10. Permeabilization solution: 0.1% (v/v) Triton X-100 diluted in PBS. 11. Fluorophore-conjugated secondary antibodies (see Note 6). 12. Coverslips mounting medium: Dissolve 1 g of Mowiol 4–88 in 4 ml PBS by gentle steering for 16 h at room temperature. Add 2 ml of anhydrous glycerol and steer for additional 16 h. Centrifuge for 15 min at 13,000  g, aliquot, and store at 4
C for several months.

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Methods

3.1 Anti-DBH Antibodies Internalization Assay

1. Plate freshly isolated chromaffin cells on fibronectin-coated coverslips at 1250 cells/mm2. Assay can be performed from 48 h to 10 days after plating (see Note 7). 2. Place cells on heated plate at 37
C. Replace cell culture medium by pre-warmed Locke’s solution. Incubate for 10 min at 37
C (see Note 8). 3. Stimulate cells for 2 min at 37
C by adding 10 μm nicotine solution in Locke’s solution. Alternatively, cells may be stimulated by a depolarizing solution of 59 mM potassium (see Subheading 2.2, item 3). 4. Transfer cells on ice and wash once with ice-cold Locke’s solution. 5. Remove Locke’s solution, and add ice-cold anti-DBH antibodies diluted in Locke’s containing 0.2% BSA. Incubate for 30 min on ice. 6. Wash cells twice with ice-cold Locke’s solution, and transfer coverslips in 37
C pre-warmed Locke’s solution to initiate endocytosis. 7. Incubate for the required time (see Note 9) before transferring back cells on ice to stop all vesicular trafficking. Maximum endocytosis is considered to be achieved after 10 min at 37
C. 8. Remove Locke’s solution, and add 4% ice-cold PFA to fix cells for 10 min at room temperature. 9. Wash three times with PBS at room temperature, and permeabilize cells for 12 min in 0.1% Triton X-100 solution. 10. Wash three times with PBS, and proceed with anti-DBH antibodies detection by adding corresponding fluorescent secondary antibodies (see Note 6) in PBS containing 3% BSA for 30 min at room temperature. 11. Wash three times with PBS, and mount coverslips in Mowiol 4–88. Store at 4
C until observation.

3.2 Image Acquisition and Analysis

1. Cells are observed under a confocal microscope. Acquisition is performed according to Nyquist parameters with high numerical aperture objectives (63, NA1.4). Pixel resolution should be around 90 nm for 488 nm excitation wavelengths, and intensity has to be carefully checked to avoid saturation. Optical section must be done at the equatorial plane of round cells to get accurate measurement of endocytosis. 2. Image files can be processed with open-source softwares like ImageJ or Icy. We will describe here the steps using Icy software which has the advantages of keeping region of interest (ROI)

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Fig. 1 Snapshot of graphical programming interface in Icy protocols. User has to define few parameters: block 1 fixes the detection threshold for spot detection, block 2 defines the image to analyze, block 4 selects the channel containing DBH staining, and blocks 6, 7, or 8 defines the scale at which spots has to be detected. Once processed, an Excel file with each spot features is generated including spot size, intensity, and EDM value which will be used to analyze spot distance from the cell edge. Value to extract can be modified in block 29 (ROI statistics). A tiff file is also generated to show the EDM and segmented spots

information in xml files associated to tiff format. In addition, Icy allows for automated image analysis without any coding knowledge, thanks to graphical programming (Fig. 1). Some features of ImageJ are embedded in Icy such as basic watershed segmentation and Euclidean distance map that are used for image analysis. Icy protocol is available on Icy website (http://icy.bioimageanalysis.org/) or upon request. 3. The first step is to delineate cells edge. Phase contrast image or, if available, cell membrane (see Note 10) or actin staining can be used. Cell delineation can be done manually (Area tool), or if staining is well contrasted, specific plug-ins can be used to find cell edges (HK-Means, Active contours). ROI are then filled in with Fill holes in ROI plug-in. 4. DBH staining is then segmented using wavelet transform decomposition of original image to efficiently detect DBH spots [8]. This is integrated in Spot detector plug-in of Icy

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Fig. 2 Quantification of anti-DBH antibodies endocytosis. (a) Bovine chromaffin cells were stimulated for 2 min with 10 μM nicotine and transferred on ice. Cells were incubated with anti-DBH antibodies for 30 min and either kept on ice for additional 5 min (stimulated, top) or returned at 37
C for 5 min (endocytosis, bottom). Before fixation, cells were incubated for 30 min with ice-cold NHS-biotin to label plasma membrane proteins. Cells were then fixed, stained with Alexa 488 fluorophore-conjugated streptavidin, permeabilized, and incubated with goat anti-rabbit antibodies conjugated to Alexa-555 fluorophore. Cells were imaged under confocal microscope and analyzed with Icy software. Cells were manually delineated using streptavidin staining and ROI (middle panel) transformed into a Euclidean distance map (EDM, right panel). Detections corresponding to DBH spots were transferred onto EDM to estimate DBH vesicles position relative to cell periphery by measuring spot mean gray intensity (right panel). (b) Distribution of DBH-positive spots (cumulative percentage of total detections per EDM values (bins of 5)). Upon endocytosis, distribution of DBH spots is shifted toward higher values. (c) Distribution of DBH-positive spots (% of total for each EDM bins). Both graphs show that 84% of detections are found at a distance of 800 nm (EDM ¼ 10) from the cell periphery in stimulated condition. After 5 min of endocytosis, this amount decreases to 32%

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(Fig. 1). Parameters have to be empirically defined and binary image compared to original image (see Note 11). 5. Since detections are exported as ROI on the original pictures, basic measurements such as mean spot intensity, spot number, and size using ROI measurements plug-in can be easily obtained. Such parameters are, for example, useful to compare exocytosis efficiency between two conditions [9, 10]. 6. To measure the efficiency of endocytosis, the position of endocytosed DBH antibodies at different incubation time at 37
C has to be defined. To do so, ROI corresponding to cell area is converted into Euclidean distance map (EDM) which gives a gray intensity to each pixel as the function of their distance to the edge of the ROI (Fig. 2a). Consequently, pixel intensity increases as it separates from the ROI edge. Then, ROI corresponding to DBH staining are transferred onto EDM picture (Fig. 2a). Mean intensity of each ROI is measured and exported into Excel files. By compiling all files, distribution of spots intensity can be obtained. An example of DBH-positive spot distribution for a single cell is shown (Fig. 2b and c). A distance threshold can be applied to estimate the percentage of stained vesicles remaining at a specific distance from the cell edge [9–11].

4

Notes 1. Coverslips are vigorously washed with detergent solution and extensively rinsed with distilled H2O. Coverslips are transferred into pure ethanol and dried out by wiping them with clean soft tissues. 2. Image analysis are performed with open-source Icy software which integrates ImageJ (http://icy.bioimageanalysis.org/). 3. Procedures to culture primary chromaffin cells from different organisms including cow, rat, and mouse have been carefully described previously [12] and in this issue [13]. 4. 0.2% (w/v) BSA diluted in Locke’s solution is used on live cells during antibody incubation to minimize nonspecific binding of antibodies. 5. Rabbit anti-dopamine-β-hydroxylase (DBH) antibodies we used was as described previously [14]. A monoclonal antiDBH antibody (Millipore, clone 4F10.2) is suitable when rabbit polyclonal antibodies are used for co-staining experiments. Each antibody lot has to be checked to adjust effective dilution. Other sources of antibody have not been tested. 6. All secondary antibodies coupled to fluorescent molecule directed against the primary antibody can be used. We

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routinely use Alexa-conjugated goat anti-rabbit antibodies at 1 μg/ml diluted in PBS containing 3% BSA. 7. Since cells tend to spread out and to adopt an elongated and flattened morphology with time in culture, we recommend performing experiments within the first 5 days of cell culture to get accurate and reproducible measurements of compensatory endocytosis. 8. Chromaffin cells in culture are highly sensitive to mechanical stress. Manipulation and changes in medium must be gentle to avoid mechanical cell stimulation and limit DBH staining in resting state. 9. After 2–3 min incubation at 37
C, most DBH staining remains close to the plasma membrane. It is, therefore, difficult to assess whether DBH is internalized or still at the cell surface. Acidic buffer might be used to strip away cell surface antibodies. Conversely, at 5 min, most DBH staining is seen inside the cell at distance from the plasma membrane. 10. For accurate cell delineation, plasma membrane proteins can be biotinylated by amine reactive biotin (Sulfo NHS-SS-Biotin) and then revealed using fluorescent streptavidin. Plasma membrane protein biotinylation can be done before or after cell fixation, but cell contour is better defined when cells are biotinylated before fixation. Following anti-DBH endocytosis assay, cells are incubated an additional 30 min on ice in PBS containing 120 μg/ml Sulfo NHS-SS-Biotin. Cells are rinsed twice in ice-cold PBS and fixed. Biotinylated proteins are revealed by incubating cells with fluorescently labeled streptavidin. 11. Typically, at 90 nm pixel resolution, sensitivity can be set to scale 2 and threshold between 30 and 70 to obtain good results. To get the best accuracy between number of spots before and after segmentation, ImageJ watershed is applied to separate spots that are too close to be resolved by spot detector. References 1. Newton AJ, Kirchhausen T, Murthy VN (2006) Inhibition of dynamin completely blocks compensatory synaptic vesicle endocytosis. Proc Natl Acad Sci U S A 103:17955–17960 2. Heuser JE, Reese TS (1973) Evidence for recycling of synaptic vesicle membrane during transmitter release at the frog neuromuscular junction. J Cell Biol 57:315–344 3. Smith C, Moser T, Xu T et al (1998) Cytosolic Ca2+ acts by two separate pathways to

modulate the supply of release-competent vesicles in chromaffin cells. Neuron 20:1243–1253 4. Artalejo CR, Elhamdani A, Palfrey HC (2002) Sustained stimulation shifts the mechanism of endocytosis from Dynamin-1-dependent rapid endocytosis to Clathrin- and Dynamin-2mediated slow endocytosis in chromaffin cells. Proc Natl Acad Sci U S A 99:6358–6363 5. Mosharov EV, Sulzer D (2005) Analysis of exocytotic events recorded by amperometry. Nat Methods 2:651–658

Compensatory Endocytosis Assay in Chromaffin Cells 6. Sankaranarayanan S, Ryan TA (2000) Realtime measurements of vesicle-SNARE recycling in synapses of the central nervous system. Nat Cell Biol 2:197–204 7. Allersma MW, Wang L, Axelrod D et al (2004) Visualization of regulated exocytosis with a granule-membrane probe using Total internal reflection microscopy. Mol Biol Cell 15:4658–4668 8. Olivo-Marin J-C (2002) Extraction of spots in biological images using multiscale products. Pattern Recogn 35:1989–1996 9. Ory S, Ceridono M, Momboisse F et al (2013) Phospholipid Scramblase-1-induced lipid reorganization regulates compensatory endocytosis in neuroendocrine cells. J Neurosci 33:3545–3556

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10. Houy S, Estay-Ahumada C, Croise´ P et al (2015) Oligophrenin-1 connects Exocytotic fusion to compensatory endocytosis in neuroendocrine cells. J Neurosci 35:11045–11055 11. Ceridono M, Ory S, Momboisse F et al (2011) Selective recapture of secretory granule components after full collapse exocytosis in neuroendocrine chromaffin cells. Traffic 12:72–88 12. Domı´nguez N, Rodrı´guez M, Machado JD et al (2012) Preparation and culture of adrenal chromaffin cells. Methods Mol Biol 846:223–234 13. Thahouly T, Tanguy E, Raherindratsara J, et al (2020) Bovine chromaffin cells: culture and fluorescent assay for secretion. This issue 14. Perrin D, Aunis D (1985) Reorganization of alpha-fodrin induced by stimulation in secretory cells. Nature 315:589–592

Chapter 4 Quantitative Methods to Study Endocytosis and Retrograde Transport of Cargo Proteins Massiullah Shafaq-Zadah, Estelle Dransart, and Ludger Johannes Abstract Endocytosis and intracellular retrograde trafficking from endosomes to the Golgi apparatus are key cellular processes. Endocytosis is directly or indirectly involved in many if not all cellular functions ranging from nutrient uptake and receptor signaling to mitosis, cell division, and migration (Scita, Di Fiore. Nature 463 (7280):464–473, 2010; McMahon, Boucrot. Nat Rev Mol Cell Biol 12(8):517–533, 2011). Retrograde trafficking is emerging as a key driver for cell polarity. Robust methods are needed to quantify these processes. At the example of the bacterial Shiga toxin and the endogenous α5β1 integrin, we here describe generic methods to differentiate (1) internalized from cell surface-accessible cargo proteins and (2) endocytic cargo proteins that have reached the Golgi apparatus via the retrograde route from those that have not. The choice of antibodies or natural ligands allows to adjust these methods to virtually any chosen biological system. Key words Endocytosis, Retrograde transport, Golgi apparatus, Covalent protein modification, Antibodies, Immunofluorescence, Immunoprecipitation, Western blot

1

Introduction In addition to its physical barrier function, the plasma membrane is the site where many active processes dynamically occur to regulate essential activities of the cell [1, 2]. Among these, protein and lipid internalization, termed endocytosis [3], is key in the regulation of many cellular functions like cell signaling, nutrients uptake, and clearance of extracellular milieu, by either unspecific fluid phase or receptor-mediated uptake [4]. Endocytosis is also required (1) during cell adhesion and migration, enabling an acute and dynamic turnover of cell adhesion molecules, such as integrins [5], and (2) during host/pathogen interaction where exogenous pathogens such as viruses or protein toxins, like the bacterial Shiga toxin, use endocytosis as an entry gate to hijack the hosts’ intracellular machinery [6].

Florence Niedergang, Nicolas Vitale and Ste´phane Gasman (eds.), Exocytosis and Endocytosis: Methods and Protocols, Methods in Molecular Biology, vol. 2233, https://doi.org/10.1007/978-1-0716-1044-2_4, © Springer Science+Business Media, LLC, part of Springer Nature 2021

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Once internalized into cells, cargo proteins are targeted to early/sorting endosomes for subsequent intracellular compartmentalization [7] to achieve specific functions. While proteins that are targeted to recycling endosomes are returned to the plasma membrane for a new round of duty, late endolysosomal compartmentalization is needed for catabolic processes, notably for clearance and nutrient uptake. Some cargoes traffic via the retrograde transport route from early/maturing endosomes to the Golgi apparatus [8]. Pathogens such as the plant toxin ricin, the bacterial Shiga or cholera toxins, and viruses such as adeno-associated virus [9] and human papillomavirus [10] all undergo retrograde transport to successively reach the Golgi apparatus and, in some cases, the endoplasmic reticulum as part of their interaction program with host cells. Endogenous proteins also undergo retrograde transport [8], but the full scope of cellular functions that is linked to this type of intracellular compartmentalization remains poorly explored. Interestingly, recent findings revealed a major contribution of retrograde transport to the polarized targeting of proteins to specialized areas of the plasma membrane [11, 12]. In this methods article, we will focus on two proteins: the exogenous receptor-binding nontoxic B-subunit of Shiga toxin (STxB) and the endogenous β1 integrin. We will provide detailed step-by-step protocols to quantitatively analyze (1) internalization, and (2) Golgi arrival by retrograde transport of these proteins, either by immunofluorescence or by immunopurification/Western blotting techniques.

2

Materials All experiments are performed in HeLa cells. The approaches can be transposed to any other cell line, however. For the retrograde transport experiments using the SNAP-tag strategy, a transgenic HeLa cell line stably expressing a Golgi-resident GalT-GFP-SNAP protein is used. For all these assays, cells are seeded 24 h prior to performing experiments in a dilution such to reach 70–80 % confluency on the day of the experiment. P4-well plates are used for immunofluorescence experiments and P6-well plates for immunopurification and Western blotting. Cells are grown at 37
C, 5 % CO2, in DMEM high-glucose glutamax medium complemented with 10 % fetal bovine serum, FBS.

2.1 General Materials

1. P4-well plates. 2. P6-well plates. 3. Glass slides. 4. 12 mm glass coverslips.

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5. Tweezers. 6. Parafilm. 7. Spin desalting columns, 7 kDa cutoff. 8. Nitrocellulose membrane. 9. Polyacrylamide gels, 4–15 % gradient. 10. Migration tank. 11. Blotting paper. 12. Transblot. 13. Chemiluminescence imaging system. 2.2 Materials for Endocytosis

1. Cy3-monofunctional maleimide reactive dye. 2. Antibodies against defined integrin conformational states: (a) 9EG7 clone (ligand-bound, “active” conformation). (b) mAb13 clone (ligand-unbound, “inactive” conformation). 3. EDTA. 4. Saponin. 5. Anti-rat IgG-Cy3 secondary antibody. 6. Mowiol: 10 % polyvinyl alcohol, 5 % glycerol, 2.5 % DABCO, 25 mM Tris buffer pH 8.5. 7. Mowiol-DAPI mounting medium: Mowiol, 1 μg/ml DAPI. 8. Acid wash buffer: 0.5 M glycine, pH 2.2, prepared in water. 9. PFA: 4 % PFA prepared in PBS. 10. PBS-BSA-SAPO: PBS, 0.2 % BSA, 0.02 % saponin. 11. PBS++ buffer: PBS complemented with 0.5 mM CaCl2 and 1 mM MgCl2. 12. NH4Cl buffer: 50 mM NH4Cl, prepared in PBS. 13. PBS-EDTA buffer: PBS, 10 mM EDTA, pH 7.4.

2.3 Material for Retrograde Transport

1. Cy3-monofunctional NHS ester reactive dye. 2. BG-maleimide, prepared in DMSO. 3. BG-GLA-NHS, prepared in DMSO. 4. DMSO anhydrous. 5. mAb13 monoclonal antibody. 6. 9EG7 monoclonal antibody. 7. SNAP-cell® block (New England Biolabs). 8. NP-40. 9. Protease inhibitor cocktail. 10. Magnetic GFP-trap beads. 11. Magnetic rack.

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12. Protein G sepharose, fast flow beads. 13. SNAP-tag® antibody (New England Biolabs). 14. Milk powder. 15. Cysteine. 16. Tween-20. 17. Anti-α tubulin antibody. 18. Anti-rabbit IgG secondary antibody. 19. Western blot protein ladders. 20. ECL femto super-signal chemiluminescence substrates. 21. Hepes buffer: 20 mM Hepes, 150 mM NaCl, pH 7.4. 22. TNE buffer: 10 mM Tris, 150 mM NaCl, 5 mM EDTA. 23. TNE lysis buffer: TNE, 1 % NP40 (V/V), 1/100 dilution of 100 protease inhibitor cocktail. 24. LSB (loading sample buffer) 3: 180 mM Tris–HCl pH 6.8, 6 % SDS, 10 % glycerol (V/V), and 0.3 mg/ml phenol red, prepared in water. 25. Migration buffer: 25 mM Tris, 250 mM glycine, 0.1 % SDS, pH 8.8. 26. Transfer buffer: 50 mM Tris, 50 mM glycine, 20 % EtOH, pH 8.8. 27. TBS buffer: 50 mM Tris, 150 mM NaCl. 28. TBS-T: TBS, 0.01 % Tween-20 (V/V). 29. TBS-T, 5 % milk: TBS-T supplemented with 5 % (W/W) milk. 30. Tris–HCl 1 M, pH 7.4.

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Methods

3.1 Endocytosis of the Exogenously Added Shiga Toxin B-Subunit (STxB) 3.1.1 Cy3 Labeling of STxB

A variant of Shiga toxin B-subunit, STxB/Cys, with an additional cysteine residue at the C-terminus of the protein was expressed and purified from bacterial E. coli as previously described [13]. Briefly, E. coli bacteria transformed with STxB/Cys encoding plasmid and grown overnight at 30
C were diluted and further incubated until reaching OD560nm ¼ 0.8. Protein expression was induced for 4 h at 42
C. Bacterial periplasm obtained after osmotic shock treatment was then submitted to a two-step fast liquid chromatography protein purification procedure, including anion exchange chromatography followed by gel filtration. 1. Dilute STxB/Cys in PBS-EDTA buffer, at a final concentration of 2 mg/ml. 2. Incubate the protein with a threefold molar excess of maleimide-Cy3 dye, for 2 h at 21
C (see Notes 1 and 2).

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3. Optional: Neutralize reaction with 10 mM free cysteine for 10 min at 21
C. 4. Eliminate the free dye by applying labeled protein on spin desalting columns 7 kDa cutoff, equilibrated with PBS, according to manufacturer’s instructions. 5. Quantify the protein concentration by measuring absorbance at 280 nm, using the extinction coefficient value of 9500 M1 cm1 for the STxB monomer. 3.1.2 STxB-Cy3 Endocytosis: “One-Wave” (i.e., Binding/Uptake) Assay

In contrast to β1 integrin for which remaining cell surface-bound antibody can be removed by acid washes (see protocol Subheading 3.2 and Note 3), no efficient method exists to remove cell surfacebound STxB. We therefore use a “one-wave” binding/uptake protocol in which STxB is bound on ice to its cellular receptor at the cell surface, the glycolipid Gb3. After washing and shift to 37
C, the protein enters cell very quickly, which thereby reduces the “background” from non-internalized STxB. Of note, all the incubations and washes are performed with 500 μl solutions. 1. For cargo binding at 4
C: Place P4 plates for 10 min on ice to block membrane dynamics, including endocytosis (see Note 4). 2. STxB-Cy3 binding: Incubate the cells with 0.4 μg/ml of STxBCy3 diluted in ice-cold complete DMEM, for 30 min at 4
C. 3. Wash the cells three times with ice-cold PBS++ buffer (see Note 5) to remove the unbound excess of STxB-Cy3 (see Note 6). For binding assay, go directly to step 5. 4. For cargo “one-wave” binding/uptake condition: Shift cells for 5 or 10 min to 37
C in complete preheated DMEM, for subsequent endocytosis of the cell surface-bound STxB-Cy3; then wash the cells three times with ice-cold PBS++ buffer. The following steps are common to surface binding (step 3) and “one-wave” binding/uptake conditions. 5. Fix the cells with ice-cold PFA for 10 min at 4
C (see Note 7). 6. Neutralize PFA with NH4Cl buffer for 10 min at RT. 7. Briefly rinse the glass coverslips in Milli-Q water. 8. Mount the glass coverslips on 7 μl Mowiol-DAPI mounting medium apposed onto glass slides, with cells facing the droplet. 9. Let dry for 30 min at 37
C.

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3.2 Endocytosis of Active and Inactive Conformations of β1 Integrin, Using an Antibody Uptake Assay 3.2.1 Anti-β1 Integrin Antibody Binding (See Note 8)

The two rat antibodies clones that are used here were raised against specific conformational states of β1 integrin: The 9EG7 antibody interacts with the active conformation, and the mAb13 antibody with the inactive one. Of note, all incubations and washes are performed with 500 μl solutions. 1. Place P4 plates for 10 min on ice to block membrane dynamics, including endocytosis (see Note 4). 2. 9EG7 or mAb13 antibody binding: Incubate the cells with 10 μg/ml of antibodies diluted in 500 μl ice-cold complete DMEM, for 30 min at 4
C (on ice). 3. Wash the cells with ice-cold PBS++ buffer to remove the unbound excess of antibodies (see Note 6). 4. Fix the cells with ice-cold PFA for 10 min at 4
C. 5. Neutralize PFA with NH4Cl buffer for 10 min at RT. 6. Saturate the cells with PBS-BSA buffer for 30 min at RT. 7. Incubate the cells with fluorophore-labeled anti-rat IgG secondary antibodies: 1:200 dilution of the secondary antibody in PBS-BSA buffer. Apply 50 μl of this solution on a parafilm. Coverslips are apposed on this droplet with tweezers, cells facing the solution. Keep in dark for 30 min at RT. 8. Wash the cells three times with PBS-BSA buffer to remove excess of unbound secondary antibodies. 9. Briefly rinse the glass coverslips in Milli-Q water. 10. Mount the glass coverslips on 7 μl Mowiol-DAPI mounting medium apposed onto glass slides, with cells facing the droplet. 11. Let dry for 30 min in a 37
C incubator.

3.2.2 Continuous Anti-β1 Integrin Antibody Uptake

1. 9EG7 or mAb13 antibody solutions: Prepare 10 μg/ml of antibodies in pre-warmed complete DMEM medium. 2. Incubate the cells with antibody solutions for 10 min at 37
C. 3. Shift the cells to 4
C (on ice) to block membrane dynamics, including endocytosis (see Note 5). 4. Wash the cells three times with ice-cold PBS++ buffer. 5. Acid wash step (see Note 3): Incubate the cells with ice-cold acid wash buffer for 45 s; remove the acid wash solution and renew it by fresh one for an additional 45 s; repeat previous step one more time (three times of 45 s incubation in total); after the last acid wash, wash the cells three times with ice-cold PBS+ + buffer. 6. Fix the cells with ice-cold PFA for 10 min at 4
C. 7. Neutralize PFA with NH4Cl buffer for 10 min at RT.

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8. Permeabilize and saturate the cells with PBS-BSA-SAPO buffer for 30 min at RT. 9. Incubate the cells with fluorophore-labeled anti-rat IgG secondary antibodies: 1:200 dilution of the secondary antibody in PBS-BSA-SAPO. Apply 50 μl of this solution on a parafilm. Coverslip are apposed on this droplet with tweezers, cells facing the solution. Keep in the dark for 30 min at RT. 10. Wash the cells three times with PBS-BSA-SAPO buffer to remove excess of unbound secondary antibodies (see Note 6). 11. Briefly rinse the glass coverslips in Milli-Q water. 12. Mount the glass coverslips on 7 μl Mowiol-DAPI mounting medium apposed onto glass slides, with cells facing the droplet. 13. Let dry for 30 min in a 37
C incubator. 3.3 Retrograde Transport of Exogenous STxB 3.3.1 Late Intracellular Delivery: Retrograde Transport of STxB-Cy3

1. Place P4 plates on ice for 10 min to block membrane dynamics, including endocytosis (see Note 4). 2. STxB-Cy3 binding: Incubate the cells with 0.4 μg/ml of STxBCy3 diluted in ice-cold complete DMEM medium, for 30 min at 4
C (on ice). 3. Wash the cells three times in ice-cold complete DMEM. 4. Shift the cells to 37
C in complete preheated DMEM for 45 min to trigger endocytic uptake. 5. Wash the cells twice with ice-cold PBS++ buffer. 6. Fix the cells with ice-cold PFA for 10 min at 4
C. 7. Neutralize PFA with NH4Cl buffer for 10 min at RT. 8. Briefly rinse the glass coverslips in Milli-Q water. 9. Mount the glass coverslips on 7 μl Mowiol-DAPI mounting medium apposed onto glass slides, with cells facing the droplet. 10. Let dry for 30 min in a 37
C incubator.

3.3.2 Benzylguanine (BG)/SNAP-tag Strategy to Study Retrograde Transport

After around 45 min of incubation with target cells at 37
C, STxB can readily be detected in Golgi membranes of HeLa cells, which are reached by retrograde transport from the plasma membrane. In contrast, the retrograde entry and exit rates of β1 integrin molecules at the level of the Golgi appear to be such that the proteins only transiently shuttles through Golgi membranes [12], making retrograde transport difficult to characterize in this case. To address this challenge, we have developed a biochemical tool to covalently “capture” retrograde cargoes that reach the Golgi, via the one-step suicide benzylguanine-SNAP reaction [14, 15]. The SNAP-tag is derived from the human DNA repair protein, O6-alkylguanine-DNA alkyltransferase. It mediates the covalent linkage of O6-benzylguanine (BG) to a specific residue of the protein, even when O6-BG is itself conjugated to other

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chemical entities [16]. Here, (1) we chemically couple our cargoes (STxB or mAb13 and 9EG7 antibodies) with the BG moiety and (2) use a transgenic cell line expressing a Golgi-localized protein, galactosyl-transferase (GalT) fused to the SNAP-tag and to GFP (termed GalT-GFP-SNAP), for biochemical pull-down or fluorescence detection. The BG-coupled cargoes that undergo retrograde transport are thereby sequestered in the Golgi apparatus, based on the covalent reaction between their BG moiety and the GalT-GFPSNAP fusion protein. BG-coupled cargoes now become detectable by immunofluorescence at the level of the Golgi apparatus and can be immunoprecipitated (GFP-trap or protein G sepharose in the case of antibody cargo). Covalent Modification of STxB/Cys with BG and Cy3

1. Dilute STxB/Cys in Hepes buffer at a final concentration of 2 mg/ml. 2. Incubate the protein overnight at 4
C with a threefold molar excess of BG-maleimide (see Notes 1 and 2). 3. Optional: Neutralize the reaction with 10 mM free cysteine for 10 min at 21
C. 4. Eliminate the free unreacted BG-maleimide using spin desalting columns 7 kDa cutoff, equilibrated with Hepes buffer according to manufacturer’s instructions. 5. Quantify the protein concentration by measuring absorbance at 280 nm, using the extinction coefficient value of 9500 M1 cm1 for the STxB monomer. 6. Incubate 200 μg of STxB-BG (at the concentration determined in step 5) in Hepes buffer for 2 h at 21
C with a threefold molar excess of NHS-Cy3 dye (see Notes 1 and 2). 7. Neutralize the reaction with 20 mM Tris for 10 min at 21
C (see Note 9). 8. Eliminate the free unreacted NHS-Cy3 dye using spin desalting columns 7 kDa cutoff, equilibrated with PBS buffer according to manufacturer’s instructions. 9. Quantify the protein concentration by measuring absorbance at 280 nm, using the extinction coefficient value of 9500 M1 cm1 for the STxB monomer.

Trapping of STxB-Cy3-BG in the Golgi: In Cellulo Localization

1. Place HeLa cells expressing GalT-GFP-SNAP seeded in P4 plates for 10 min on ice to block membrane dynamics, including endocytosis (see Note 4). 2. STxB-Cy3 +/BG (BG-modified or non-modified STxB-Cy3) binding: Incubate the cells with 0.4 μg/ml of STxB-Cy3 +/ BG for 30 min at 4
C, in ice-cold complete DMEM medium. 3. Wash the cells three times in ice-cold complete DMEM. 4. Shift the cells to 37
C for 4 h in complete preheated DMEM medium.

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5. Wash the cells twice with ice-cold PBS++ buffer. 6. Fix the cells with ice-cold PFA for 10 min at 4
C. 7. Neutralize PFA with NH4Cl buffer for 10 min at RT. 8. Briefly rinse the glass coverslips in Milli-Q water. 9. Mount the glass coverslips on 7 μl Mowiol-DAPI mounting medium apposed onto glass slides, with cells facing the droplet. 10. Let dry for 30 min at 37
C. Trapping of STxB-Cy3-BG in the Golgi: Biochemical Characterization

In all these experiments, incubations and washes are done with 1 ml solutions, except the lysis step which is done with 0.5 ml solution. 1. Place HeLa cells expressing GalT-GFP-SNAP seeded in P6 plates for 10 min on ice to block membrane dynamics, including endocytosis (see Note 4). 2. STxB-Cy3 +/BG binding: Incubate the cells with 0.4 μg/ml of STxB-Cy3 +/BG for 30 min at 4
C (on ice), diluted in ice-cold complete DMEM medium. 3. Wash the cells in ice-cold complete DMEM. 4. Shift the cells to 37
C for 4 h in complete preheated DMEM medium. 5. Discard medium and incubate cells with SNAP-cell block solution for 20 min at 37
C, diluted (1/200) in complete preheated DMEM medium (see Note 10). 6. Wash the cells twice with ice-cold PBS++ buffer. 7. Lyse the cells with 500 μl TNE lysis buffer for 30 min on ice. 8. Scrape the cells off the wells with 1 ml tips whose ends have been cut, and transfer them in 1.5 ml tubes. 9. Centrifuge the cells at maximal speed for 10 min at 4
C (PNS: postnuclear supernatant). 10. In parallel, for each condition, prepare 30 μl GFP-trap beads in 1.5 ml tubes as follows (see Note 11). 11. Wash the beads in 500 μl TNE buffer. 12. Discard the supernatant using a magnetic rack. 13. Repeat steps 11 and 12 two more times. 14. After the last wash, remove as much liquid as possible. 15. Apply the PNS from step 9 to the prepared beads. 16. Incubate overnight at 4
C on a rotating wheel. 17. Save 30 μl of the unbound fraction. Remove and discard the remaining supernatant and wash the beads as described in step 10. 18. Resuspend the dried beads (using a Hamilton syringe) with 40 μl of LSB 1.5 and add 15 μl LSB 3 to the 30 μl of each unbound fraction.

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19. Boil the beads and the unbound fractions for 10 min at 95
C to elute and denature proteins. 20. SDS-PAGE analysis: Load the samples on two polyacrylamide 4–15 % gradient gels, gel 1 for the eluted proteins, and gel 2 for the unbound fractions. Load 5 μl of Western blot protein ladders for protein size standard and run at 120 V for 10 min, then 200 V until the end of migration. 21. Equilibrate the gels, the nitrocellulose membranes, and the blotting papers in transfer buffer. 22. Transfer proteins (semidry transblot) on nitrocellulose membranes for 1 h at 10 V. 23. Incubate the membranes in TBS-T, 5 % milk buffer, for 1 h at room temperature (RT). 24. Incubate the membrane 1 with primary antibody against STxB and the membrane 2 with anti α-Tubulin, for 2 h at RT (dilution in TBS-T, 5 % milk buffer, according to manufacturer’s instructions). 25. Wash the membranes with TBS-T buffer three times for 5 min at RT (see Note 6). 26. Incubate the membranes 1 and 2 with HRP-coupled secondary anti-rabbit or anti-mouse IgG antibodies, respectively (1/5000 dilution in TBS-T, 5 % milk buffer), for 45 min at RT. 27. Wash the membranes with TBS-T buffer three times for 5 min at RT (see Note 6). 28. Incubate the membranes with HRP substrate (Femto ECL). 29. Acquire the chemiluminescence signal. 3.4 Retrograde Transport of Active and Inactive Conformation-Specific Anti-β1 Integrin Antibodies Using the BG/SNAP Approach 3.4.1 BG Labeling of β1 Integrin Antibodies (mAb13 and 9EG7) 3.4.2 Trapping of Anti-β1 Integrin Antibodies in the Golgi: Biochemical Characterization

1. Use antibodies at a final concentration of 0.5–1 mg/ml. 2. Incubate the protein with a tenfold molar excess of BG-GLANHS for 6 h at 4
C. 3. Neutralize the reaction with 20 mM Tris for 10 min at 21
C (see Note 9). 4. Eliminate the free BG-GLA-NHS by applying modified antibodies on spin desalting columns 7 kDa cutoff, equilibrated with PBS, according to manufacturer’s instructions.

In these experiments, we track retrograde β1 integrin trafficking using specific antibodies coupled to BG. This approach is termed antibody uptake experiment and is based on the following idea: If β1 integrin itself undergoes retrograde transport to the Golgi apparatus, BG-tagged anti-β1 integrin antibodies that are bound to β1

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integrin at the cell surface should follow the same fate and be trapped in this compartment via the BG-SNAP reaction. Of note, in all these experiments, incubations and washes are done with 1 ml solutions, except the lysis step which is done with 0.5 ml solutions. 1. Incubate GalT-GFP-SNAP-expressing HeLa cells seeded in P6 plates for 4 h at 37
C with 10 μg/ml of mAb13 or 9EG7, BG modified or not (i.e., mAb13, mAb13-BG, 9EG7, 9EG7-BG). 2. Discard the medium, and neutralize the unreacted SNAP with a 1/200 dilution of SNAP-cell block solution for 20 min at 37
C (see Note 10). 3. Wash the cells three times with ice-cold PBS++ buffer. 4. Lyse the cells with 500 μl TNE lysis buffer for 30 min on ice. 5. Scrape the cells off the wells with 1 ml tips whose ends have been cut, and transfer to 1.5 ml tubes. 6. Centrifuge the cell lysates at maximal speed for 10 min at 4
C (PNS: post-nuclear supernatant). 7. In parallel, for each condition, prepare 50 μl protein G sepharose beads in 1.5 ml tubes (see Note 11) as follows. 8. Wash the beads in 500 μl TNE buffer. 9. Centrifuge at 1500  g for 5 min. 10. Discard supernatant. 11. Repeat the washes two more times. 12. After the last wash, remove as much liquid as possible using a Hamilton syringe. 13. Apply the PNS from step 6 to the prepared beads. 14. Incubate overnight at 4
C on a rotating wheel. 15. Centrifuge at 1500  g for 5 min; save 30 μl of the unbound fraction and discard the remaining supernatant; wash the beads as described from step 9 to 11. 16. After the last wash, remove as much liquid as possible from the beads using a Hamilton syringe. 17. Resuspend the dried beads with 40 μl LSB 1.5. 18. Boil the beads and the unbound fractions for 10 min at 95
C to elute and denature proteins. 19. SDS-PAGE analysis: Load the samples on two polyacrylamide 4–15 % gradient gels, gel 1 for the eluted proteins, and gel 2 for the unbound fractions. Load 5 μl of Western blot protein ladders for protein size standards and run for 10 min at 120 V and then 200 V until the end of migration. 20. Equilibrate the gels, the nitrocellulose membranes, and the blotting papers in transfer buffer.

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21. Transfer the proteins (semi-dry transblot) for 1 h at 10 V onto nitrocellulose membranes. 22. Incubate the membranes in TBS-T, 5 % milk solution, for 1 h at RT on a gentle rocker shaker. 23. Cut the milk-saturated membrane 1 under the 250 kDa molecular weight marker, keep the upper one (immunoreactive band of interest at around 280 kDa). Cut the milk-saturated membrane 2 above the 70 kDa molecular weight marker, keep the lower one (immunoreactive band of interest at around 55 kDa). The two resulting membranes are further processed for 2 h at RT, as follows. 24. Incubate the upper part of the membrane 1 with primary antibody against SNAP-tag protein (1/1000 dilution in TBS-T, 5 % milk solution) for the detection of β1 integrin antibody-GalT-GFP-SNAP product (280 kDa). 25. Incubate the lower part of the membrane 2 with primary antibody against α-tubulin (dilution in TBS-T, 5 % milk solution, according to manufacturer’s instructions) as loading control (55 kDa) (see Note 12). 26. Wash the membranes three times with TBS-T buffer for 5 min at RT on a strong rocker shaker (see Note 6). 27. Incubate the membranes for 45 min at RT with anti-rabbit IgG (SNAP) or anti-mouse IgG (tubulin), HRP-coupled secondary antibody, 1/5000 dilution in TBS-T, 5 % milk solution. 28. Wash the membranes three times with TBS-T buffer for 5 min at RT (see Note 6). 29. Incubate the membranes with HRP substrate (Femto ECL). 30. Acquire the chemiluminescence signal. 3.5 Quantification Using ImageJ Software (https:// imagej.nih.gov/ij/) 3.5.1 Quantification of Binding and Endocytosis Processes Membrane-Bound Fluorescent Signal: mAb13 and 9EG7 Internalized Fluorescent Signal: mAb13 and 9EG7 Endocytosis

1. Use the “Freehand selections” item in ImageJ program to delimit the contour of cells (white dashed line). 2. Go to “Analyze” and “Measure” (Fig. 1a). A window displaying different parameters will appear: the area, the mean intensity, and the integrated density (IntDen), among others. 3. For this quantification, we use the “mean” intensity measurement. For each experimental condition, this operation is typically repeated on indicated number of cells per experiment. Perform three or more independent experiments to finally get an average mean intensity (see Note 13). We use a similar methodology (Fig. 1b). Since the acid wash step efficiently removes the membrane-bound pool of ligand (except for STxB), the difficulty here is to accurately delimit the cell boundary.

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Fig. 1 Anti-β1 integrin antibodies binding and endocytosis assay. (a) Binding: 10 μg/ml of mAb13 or 9EG7 antibodies were incubated for 30 min at 4
C with HeLa cells. Cells were washed, fixed, and immuno-stained with fluorescent-labeled secondary antibodies. As expected, only membrane staining is observed. Of note, the labeling with mAb13 is stronger than that with 9EG7. (b) Endocytosis: Cells were incubated for 10 min at 37
C in the continuous presence of 10 μg/ml of mAb13 or 9EG7 antibodies. The remaining cell surface-accessible antibodies were removed by acid wash, prior to fixation. Cells were then permeabilized with saponin (PBS-BSA-SAPO buffer) before being processed as in (a). (c) Same as (b), except that the signal was artificially enhanced to visualize the contour of the cell. (d) Endocytosis quantification: The internalized fractions of mAb13 (n ¼ 32 cells) and 9EG7 (n ¼ 32 cells) were quantified using the ImageJ program. All images were acquired on a confocal microscopy. White dashed lines delimit cell boundaries. Scale bars, 10 μm

For cell delimitation (but not for quantification), we therefore artificially push the signal to the maximum level in a way such that the background is highly enhanced, thus allowing the approximative visualization of the whole cell (Fig. 1c). Quantification is shown in Fig. 1d (see Note 13). To compare the endocytosis of a given cargo (the inactive conformation of β1 integrin labeled by mAb13, for instance) in different experimental conditions (knockdown, small molecule inhibitor treatments, agonist stimulation, etc.), it is mandatory to

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Fig. 2 Retrograde transport of STxB or of conformational state-specific anti-β1 integrin antibodies. (a) Schematic representation of a cell with the following features used for the quantification methodology: the perinuclear Golgi compartment (white dashed line), the cell boundary (pink dashed line), and the nucleus (blue). (b) Detection of STxB retrograde transport by immunofluorescence: HeLa cells stably expressing the GalT-GFP-SNAP fusion protein were incubated at 37
C with STxB-Cy3 for 45 min, with STxB-Cy3 for 4 h, or with STxB-Cy3-BG for 4 h, in one-wave binding uptake experiments. (c) The percentage of Golgi-localized STxB was quantified from experiments in (b) on 10–15 cells per condition. Note that after 4 h on incubation, STxB-Cy3-BG shows a more efficient Golgi retention when compared to the unmodified STxB-Cy3. (d, e) Retrograde transport detection by pull-down experiments and Western blotting: The covalent reaction products between STxB-BG or BG-coupled conformational state-specific anti-β1 integrin antibodies and Golgi-localized GalT-GFP-SNAP fusion protein were purified using GFP-trap or protein G sepharose-coated beads. The elutes of denatured proteins from these beads (LSB boiled at 95
C) were further analyzed by immunoblotting (IB) against STxB (IB: anti-STxB) or SNAP-tag (IB: anti-SNAP), respectively. For protein normalization, α-Tubulin immunoblotting was performed. All images are acquired by confocal microscopy. Scale bars, 10 μm

normalize the endocytosis measurement with the initial binding efficiency of this cargo in all conditions to avoid any bias in the final quantification. Thus, if any variation is occurring at the binding step (e.g., if a treatment modality affects the total amount of a given receptor that is present at the plasma membrane at steady state), this will be considered for the quantification of endocytosis rates.

Endocytosis and Retrograde 3.5.2 Quantification of Cargo Accumulation within the Golgi Compartment Percentage of Golgi-Localized Cargo Using Immunofluorescence: STxB-Cy3 or STxB-Cy3-BG

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After a 45 min one-wave uptake, STxB clearly accumulates in the Golgi (Fig. 2b, left panel). After longer times of incubation, such as 4 h in our experiment, the majority of STxB has exited this compartment and is now localized in the endoplasmic reticulum (Fig. 2b, middle panel). In contrast, in the condition where the BG/SNAP-tag strategy is used, STxB-BG is now retained in the Golgi compartment, even at the 4 h time point, which is due to its covalent reaction with the Golgi-resident GalT-GFP-SNAP fusion protein (Fig. 2b, right panel). For the quantification, we measure two parameters: 1. Golgi signal of STxB-Cy3 or STxB-Cy3-BG: Use the “Freehand selections” tool of ImageJ (see above) to delimit the Golgi apparatus (Fig. 2a, scheme, white dashed line) based on the GFP channel (i.e., GalT-GFP-SNAP: Golgi marker). Then switch to Cy3 channel (STxB-Cy3), and click on “Analyze” and then “Measure” tabs. 2. Total signal: Use the “Freehand selection” to delimit the whole cell area (Fig. 2a, scheme, pink dashed line), and click on “Analyze” and then “Measure” tabs. In contrast to the previous measurements for endocytosis, the integrated density “IntDen” is used instead of the mean value. The integrated density corresponds to the product of the mean value and the area. Indeed, area sizes of Golgi and whole cell vary considerably. The sum of pixel values within the selected areas is therefore quantified. The final Golgi ratio measurement is the following: (Golgi intensity/total cell intensity)  100 (Fig. 2c).

Relative Amount of Golgi-Retrieved Protein (mAb13, 9EG7, or STxB, +/ BG): Immunopurification and Western Blotting

BG-modified cargoes covalently react with GalT-GFP-SNAP if they are retrieved from the plasma membrane to the Golgi apparatus. The resulting reaction products are immunopurified and detected by Western blotting (Fig. 2d, e). Note that when BG-modified ligands are used, GalT-GFP-SNAP-STxB and GalT-GFP-SNAPmAb13 are detected, in contrast to 9EG7. 1. For quantification, we use the “Rectangle” tool to delimit the protein bands in the different experimental conditions (red rectangle). 2. Click on the following tabs: “Analyze,” “Gel,” “select first lane,” then back to “Analyze,” “Gel,” and “Plot lanes” (Fig. 2e, plot profile). At this level, it is important to close each individual plot area, corresponding to the protein signal density of each condition, using the “Straight line” tool (Fig. 2d, e, plot profile, red lines). 3. Use the “wand (tracing) tool,” and click onto each individual plot to get the intensity value of the protein bands (Fig. 2d, e,

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graphs). For normalization of the signal to the total amount of initial material, we use tubulin (“housekeeping protein”) immunoblot detection. The amount of Golgi-retrieved protein is then quantified based on the following ratio: (GalT-GFP-SNAP-cargo band intensity)/(α-tubulin band intensity).

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Notes 1. For the covalent modification of STxB, the molar excess of dyes or BG is calculated considering the molecular weight of the STxB monomer. 2. When coupling proteins (STxB or β1 integrin antibodies) with BG or Cy3, using NHS ester or maleimide chemistry, reaction buffers should be devoid of any preservatives such as gelatin, BSA, or any other protein content. The presence of glycerol may also decrease coupling efficiency. 3. Acid washes are an important step to remove cell surfaceexposed β1 integrin antibodies, such that “endocytosis” quantification can be performed in a meaningful manner. Do not exceed indicated acid wash incubation times to preserve cellular integrity. 4. For protein binding experiment (STxB or β1 integrin antibodies), performing experiments strictly at 4
C on ice with buffers precooled and kept at 4
C is mandatory to avoid any premature internalization of the cargoes. 5. Using PBS++ buffer rather than PBS is recommended for the washes steps in order to minimize the detachment of cells from glass coverslips or plastic dishes. 6. Efficient washing procedures are of prime importance for immunofluorescence and Western blotting to efficiently decrease background signal that may affect final quantifications. 7. PFA should be handled carefully under the fume hood. 8. The “binding assay” is necessary to normalize endocytosis measurements, in case variations occur at the binding step due to experimental conditions (e.g., decreased or increased molecular quantities of receptors at the cell surface). Such variations in total cell-associated ligands would bias the quantification of the uptake step. 9. NHS ester chemistry efficiently targets primary amines, notably on lysines. NHS is quenched by the primary amines of Tris buffer.

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10. Prior to cell lysis, it is critical to neutralize unreacted SNAP-tag using SNAP block reagent to avoid post-lysis reactions. 11. To immunopurify BG-SNAP reaction products between STxB or β1 integrin antibodies and Golgi-localized GalT-GFPSNAP, GFP-trap or protein G sepharose beads are used, respectively. 12. As in the case of the “binding assay,” tubulin is used to normalize the Western blot signals from covalent reaction products between STxB or β1 integrin antibodies and GalTGFP-SNAP to the total protein amount of cell lysate. 13. During signal acquisition at the confocal microscope and for further imaging processing with ImageJ, make sure to apply strictly the same settings in all conditions. Otherwise, quantifications will not be meaningful. References 1. Scita G, Di Fiore PP (2010) The endocytic matrix. Nature 463(7280):464–473. https:// doi.org/10.1038/nature08910 2. McMahon HT, Boucrot E (2011) Molecular mechanism and physiological functions of clathrin-mediated endocytosis. Nat Rev Mol Cell Biol 12(8):517–533. https://doi.org/10. 1038/nrm3151 3. Kumari S, Mg S, Mayor S (2010) Endocytosis unplugged: multiple ways to enter the cell. Cell Res 20(3):256–275. https://doi.org/10. 1038/cr.2010.19 4. Doherty GJ, McMahon HT (2009) Mechanisms of endocytosis. Annu Rev Biochem 78:857–902. https://doi.org/10.1146/ annurev.biochem.78.081307.110540 5. Moreno-Layseca P, Icha J, Hamidi H, Ivaska J (2019) Integrin trafficking in cells and tissues. Nat Cell Biol 21(2):122–132. https://doi. org/10.1038/s41556-018-0223-z 6. Sandvig K, Grimmer S, Lauvrak SU, Torgersen ML, Skretting G, van Deurs B, Iversen TG (2002) Pathways followed by ricin and Shiga toxin into cells. Histochem Cell Biol 117 (2):131–141. https://doi.org/10.1007/ s00418-001-0346-2 7. Jovic M, Sharma M, Rahajeng J, Caplan S (2010) The early endosome: a busy sorting station for proteins at the crossroads. Histol Histopathol 25(1):99–112. https://doi.org/ 10.14670/HH-25.99 8. Johannes L, Popoff V (2008) Tracing the retrograde route in protein trafficking. Cell 135 (7):1175–1187. https://doi.org/10.1016/j. cell.2008.12.009

9. Nonnenmacher ME, Cintrat JC, Gillet D, Weber T (2015) Syntaxin 5-dependent retrograde transport to the trans-Golgi network is required for adeno-associated virus transduction. J Virol 89(3):1673–1687. https://doi. org/10.1128/JVI.02520-14 10. Schelhaas M, Ewers H, Rajamaki ML, Day PM, Schiller JT, Helenius A (2008) Human papillomavirus type 16 entry: retrograde cell surface transport along actin-rich protrusions. PLoS Pathog 4(9):e1000148. https://doi.org/10. 1371/journal.ppat.1000148 11. Carpier JM, Zucchetti AE, Bataille L, Dogniaux S, Shafaq-Zadah M, Bardin S, Lucchino M, Maurin M, Joannas LD, Magalhaes JG, Johannes L, Galli T, Goud B, Hivroz C (2018) Rab6-dependent retrograde traffic of LAT controls immune synapse formation and T cell activation. J Exp Med 215(4):1245–1265. https://doi.org/10.1084/jem.20162042 12. Shafaq-Zadah M, Gomes-Santos CS, Bardin S, Maiuri P, Maurin M, Iranzo J, Gautreau A, Lamaze C, Caswell P, Goud B, Johannes L (2016) Persistent cell migration and adhesion rely on retrograde transport of beta(1) integrin. Nat Cell Biol 18(1):54–64. https://doi.org/ 10.1038/ncb3287 13. Su GF, Brahmbhatt HN, Wehland J, Rohde M, Timmis KN (1992) Construction of stable LamB-Shiga toxin B subunit hybrids: analysis of expression in salmonella typhimurium aroA strains and stimulation of B subunit-specific mucosal and serum antibody responses. Infect Immun 60(8):3345–3359 14. Shi G, Azoulay M, Dingli F, Lamaze C, Loew D, Florent JC, Johannes L (2012)

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SNAP-tag based proteomics approach for the study of the retrograde route. Traffic 13 (7):914–925. https://doi.org/10.1111/j. 1600-0854.2012.01357.x 15. Johannes L, Shafaq-Zadah M (2013) SNAPtagging the retrograde route. Methods Cell Biol 118:139–155. https://doi.org/10. 1016/B978-0-12-417164-0.00009-4

16. Keppler A, Gendreizig S, Gronemeyer T, Pick H, Vogel H, Johnsson K (2003) A general method for the covalent labeling of fusion proteins with small molecules in vivo. Nat Biotechnol 21(1):86–89. https://doi.org/10.1038/ nbt765

Chapter 5 High-Content Drug Discovery Screening of Endocytosis Pathways David A. Cardoso, Ngoc Chau, and Phillip J. Robinson Abstract Endocytosis is the dynamic internalization of cargo (receptors, hormones, viruses) for cellular signaling or processing. It involves multiple mechanisms, classified depending on critical proteins involved, speed, morphology of the derived intracellular vesicles, or substance trafficked. Pharmacological targeting of specific endocytosis pathways has a proven utility for diverse clinical applications from epilepsy to cancer. A multiplexable, high-content screening assay has been designed and implemented to assess various forms of endocytic trafficking and the associated impact of potential small molecule modulators. The applications of this assay include (1) drug discovery in the search for specific, cell-permeable endocytosis pathway inhibitors (and associated analogues from structure-activity relationship studies), (2) deciphering the mechanism of internalization for a novel ligand (using pathway-specific inhibitors), (3) assessment of the importance of specific proteins in the trafficking process (using CRISPR-Cas9 technology, siRNA treatment, or transfection), and (4) identifying whether endocytosis inhibition is an off-target for novel compounds designed for alternative purposes. We describe this method in detail and provide a range of troubleshooting options and alternatives to modify the protocol for lab-specific applications. Key words Endocytosis, Pharmacology, Therapeutics, Drug discovery, High-content screening

Abbreviations CLIC/GEEC CME DAPI DMEM DMSO EGFR FBS FEME GFP IXM PBS PFA

clathrin-independent carriers/glycosylphosphatidylinositol-anchored proteinenriched endocytic compartments clathrin-mediated endocytosis 40 -6-diamidino-2-phenylindole Dulbecco’s Modified Eagle Medium dimethyl sulfoxide epidermal growth factor receptor fetal bovine serum fast endophilin-mediated endocytosis green fluorescent protein ImageXpress Micro phosphate-buffered saline paraformaldehyde

Florence Niedergang, Nicolas Vitale and Ste´phane Gasman (eds.), Exocytosis and Endocytosis: Methods and Protocols, Methods in Molecular Biology, vol. 2233, https://doi.org/10.1007/978-1-0716-1044-2_5, © Springer Science+Business Media, LLC, part of Springer Nature 2021

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Introduction Endocytosis broadly refers to the active transport of extracellular ligands into the cytoplasm for intracellular processing via invaginations of the lipid plasma membrane [1]. Coupled with exocytosis, which is the directionally opposing route of membrane trafficking, a dynamic exchange of information between cells and their immediate environment can be established. Various physiological outcomes can be achieved through the internalization of cargo, including but not limited to nutrient intake (vitamins, ironbound transferrin; cell health), cell-surface receptor activation or modulation (tyrosine kinases; signal transduction, desensitization), and particle entry (viruses such as HIV; infection). Each of these instances requires the active functioning of at least one endocytosis pathway. The endocytic modes are characterized and subdivided on numerous functional and structural characteristics such as the more critical or unique proteins involved, rapidity of internalization, and the morphology of the resultant vesicle or cargo trafficked [2, 3]. Several ubiquitous, well-characterized endocytic pathways have been identified that are suitable for high-content analysis: (1) clathrin-mediated endocytosis (CME), (2) fast endophilinmediated endocytosis (FEME), (3) caveolae-mediated endocytosis, (4) clathrin-independent carriers (CLIC)/glycosylphosphatidylinositol-anchored protein-enriched endocytic compartments (GEEC) pathway, (5) arf6-associated endocytosis, and (6) rho-dependent IL-2 receptor endocytosis (Fig. 1). There are also additional pathways such as fluid phase endocytosis (macropinocytosis) and phagocytosis. Particular ligands have been identified which are preferentially internalized by specific pathways; for example, transferrin and epidermal growth factor receptors (EGFR) are internalized by CME. However, at high EGF concentrations, the EGFR traffics by alternative pathways such as FEME. Thus, it is uncommon for cargo to be wholly selective to a single pathway under all conditions. Pharmacological targeting of endocytic pathways is an important avenue for discovery and development of several potential therapeutics. There are various proof-of-concept studies for the efficacy of endocytosis modulators in epilepsy, cancer, or viral and bacterial infections [4–6]. Epilepsy is a debilitating disorder characterized by the hyper-synchronized and uncontrolled firing of neurons, culminating in seizure activity. The continuous neuronal activity can continue due to the recycling/replenishment of vesicles (for neurotransmitter reuptake into nerve terminals and for later release into the synaptic cleft) via ultrafast endocytosis, CME, and activity-dependent bulk endocytosis, all of which are dynamindependent. Inhibition of dynamin, a GTPase mechanoenzyme

Fig. 1 Endocytosis pathways suitable for analysis using this protocol; highlighting vesicle morphology, key proteins involved, preferentially internalized cargo, and known small molecule inhibitors. (a) CME. (b) FEME. (c) Caveolae-dependent endocytosis. (d) CLIC/GEEC pathway. (e) Arf6-associated endocytosis. (f) Rho-dependent IL-2 endocytosis. Not shown are additional pathways such as fluid phase endocytosis (macropinocytosis) or phagocytosis

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responsible for the rate-limiting scission step of many forms of endocytosis, by small molecules such as dynasore (a less potent analogue of dyngo®-4a), has shown effectiveness in limiting the propagation of seizures, presumably by limiting the recycling of synaptic vesicles [5]. Infectious particles (e.g. bacterial toxins or HIV) are capable of hijacking host cellular machinery and causing disease via internalization through specific endocytic pathways. Pharmacological intervention has been proposed as a viable avenue for prophylaxis against diseases resulting from certain pathogenic bacteria, viruses, or toxins [4] as demonstrated for botulism in mice [7]. Suppressing endocytosis of EGFR is a promising approach to lung cancer [8]. With these examples, it is evident that inhibition of endocytosis has broad utility in discovery and development of a variety of therapeutics. Herein, we report an adaptable, multiplexable, high-content endocytosis screening assay capable of identifying chemical modulators (inhibitors or activators) of specific endocytic pathways, as well as establishing the importance of certain proteins in a given endocytic route. The protocol focuses primarily on screening for novel inhibitors of transferrin-mediated CME using the humanderived U2OS cell line, with points of adaptation to other endocytic modes and optimization being noted throughout the text.

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Materials and Solutions

2.1 Cell Cultivation and Seeding

Dulbecco’s phosphate-buffered saline (PBS), Dulbecco’s Modified Eagle Medium (DMEM) high glucose, fetal bovine serum (FBS) qualified, trypsin-EDTA solution, fibronectin from bovine plasma, tissue culture flask vented (25 cm2, 75 cm2, and 150 cm2), Falcon® conical polypropylene tubes (15 mL and 50 mL), Costar® Stripette® pipettes (5 mL, 10 mL, and 25 mL), Nalgene™ RapidFlow™ filter unit (500 mL, 0.2 μm), MatriPlate™ 96-well glassbottom micro-well plate (Brooks) (MGB096-1-2-LG-L).

2.2

Powdered compound stock (e.g. dyngo®-4a, dynamin inhibitor (Abcam) (ab120689); pitstop®-2, clathrin inhibitor (Abcam) (ab120687); amiloride, Na+/H+ pump inhibitor), dimethyl sulfoxide (DMSO), polypropylene 96-well V-bottom microplate (Greiner Bio-One) (651201), bovine serum albumin, Tween-80.

Pharmacology

2.3 Endocytosis Assay

Paraformaldehyde (PFA) extra pure, sodium chloride, acetic acid (glacial), Alexa Fluor™-594 transferrin (5 mg), 40 -6-diamidino-2phenylindole (DAPI; 5 mg).

2.4 Equipment and Software

Ovation E12 automated 5–250 μL and 25–850 μL pipettes (VistaLab Technologies), ImageXpress Micro (IXM) XLS Widefield High-Content Analysis System (MetaMorph software) (Molecular Devices), GraphPad Prism (GraphPad Software Incorporated).

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Solutions

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Cell culture media (500 mL): DMEM supplemented with 10% FBS. 1. Transfer 50 mL of DMEM from the 500 mL stock bottle into a sterile Falcon tube (can be used in the cell serum starvation step). 2. Add 50 mL of heat-inactivated FBS to remaining DMEM. 3. Filter sterilize through suction filter unit (0.2 μm), and store at 4  C for repeat uses.

2.5.1 Blocking Solution (500 mL): 0.5% V/V Tween-80 and 0.1% W/V Bovine Serum Albumin

1. Dissolve 0.5 g bovine serum albumin and 2.5 mL Tween-80 into 300 mL MQ water.

2.5.2 Low-pH Acid Wash (500 mL): 0.2 M Acetic Acid and 0.5 M NaCl, pH 2.8

1. Dissolve 14.6 g NaCl and 5.8 mL acetic acid (glacial) in 300 mL MQ water.

2.5.3 PFA (4%) in PBS (500 mL)

1. Dissolve 20 g of PFA powder into 250 mL pre-warmed PBS (60  C).

2. Make up remaining volume in MQ water, filter, and store at 4  C until use.

2. Make up the remaining volume in MQ water, adjust pH to 2.8 with NaOH solution, filter (0.22 μm), and store at 4  C until use.

2. Add 2–3 drops of 10 M NaOH to promote dissolution of PFA. 3. Adjust pH to 7.4, filter sterilize (0.22 μm), and aliquot out into 50 mL portions. 4. Store at 20  C and thaw when required. If PFA is cloudy upon thawing, the solution was prepared improperly (e.g. incorrect pH) and thus cannot be used. 2.5.4 DAPI Stain (200 μg/ mL)

1. Add 1 mL of pure methanol to dissolve DAPI powder.

2.5.5 Alexa Fluor™-594 Transferrin (5 μg/μL)

1. Resuspend transferrin powder in 1 mL of MQ water, and store at 4  C until required (minimizing light exposure).

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2. Add 24 mL of MQ water, aliquot out into 500 μL portions, and store at 4  C.

Method

3.1 Preparation of Cells for Endocytosis Assay

1. Cell stock revival and culture: Using aseptic tissue culture techniques, revive cell type of choice (U2OS, COS7 or HeLa cells) from liquid N2 frozen stock and upscale with 1–2 passages to achieve the required cell number for the planned experiment. For example, a T150 flask of U2OS cells is sufficient for three 96-well plates or twelve test compound conditions (see

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Note 1). Herein, “media” refers to the cell culture media suitable for a particular cell line (DMEM + 10% FBS for U2OS/COS7 cells; RPMI-1640 + 10% FBS for HeLa cells). The cell type can also include those that have been engineered (e.g. using CRISPR-Cas9) to have a permanent or conditional knockout of a protein of interest to assess its impact on endocytic pathways. 2. Cell preparation (optional): Prior to seeding, transfection (green fluorescent protein (GFP)-tagged protein plasmid overexpression or siRNA-mediated knockdown) may be required. Perform as required and allow expression/knockdown to take effect prior to initiating next step of the protocol. Day 1: Seeding cells 3. Plate preparation: Coat MatriPlate™ 96-well glass-bottom plate with 60–100 μL/well of 50 μg/mL fibronectin solution using a multichannel pipette (25–850 μL) to promote cell adhesion to the glass surface. Fresh fibronectin solution (50 μg/mL) is made by diluting 1 mg/mL fibronectin stock (1:20) with pre-warmed PBS. It is important to ensure that the solution covers the entire surface of each well. This can be achieved by gently tapping the side of the plate to disperse the solution. Incubate the plate at 37  C for at least 2 h prior to seeding cells. 4. Cell seeding: Trypsinize and pellet the cells using a benchtop centrifuge (190  g, 3–5 min). Aspirate the supernatant and resuspend the cell pellet in 3–5 mL media (depending on the cell pellet size). Determine cell concentration (number of cells/mL) using a hemocytometer (or equivalent method). Referring to Example Calculation, determine and perform the appropriate dilution of the cell suspension required for plate seeding. Following time elapse, flick off the fibronectin solution, and wash the plate(s) three times with 200 μL/well warm PBS using a multichannel pipette (25–850 μL). Proceed to pipette 200 μL/well of diluted cell solution using the lowest pipette speed (to avoid sheer stress). Allow the plate(s) to sit on the bench at room temperature for 10–15 min to ensure all cells settle to the bottom of the plate(s) and then transfer to an incubator and leave overnight (37  C, 5% CO2).

Example Calculation For U2OS cells, a seeding density of 7000–9000 cells per well allows for the achievement of a monolayer of cells in wells that is required for high-content imaging and analysis.

(continued)

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Hemocytometer ¼ 62 cells counted per chamber (for example). Cell stock solution ¼ 62  104 cells/mL ¼ 620,000 cells/mL. Seeding density ¼ 7000 cells/well (for example). Seeding volume ¼ 200 μL/well. Dilution factor ¼ (620,000  5)  7000 ¼ 17.7. Volume required per plate: 200 μL/well  96 wells ¼ 19.2 mL. Make 25 mL diluted cell solution ¼ 1.4 mL concentrated cell solution + 23.6 mL pre-warmed media.

Day 2: Serum starvation and endocytosis assay 5. Serum starvation: To limit cell division, promote cellular adherence, and to maximize uptake of transferrin, serum starvation (from 2 h to overnight) is performed (i.e. removal of FBS from the media). Remove cell plates from the incubator, and confirm that cells have attached and spread evenly on the fibronectin-coated well surface, using a light microscope (see Note 2). Flick off media to waste and replace with 200 μL/well warm (37  C) media without FBS. For more sensitive cell lines, use serum-reduced Opti-MEM media. Place plate(s) back into the incubator (37  C, 5% CO2), and incubate for 2 h to overnight (see Note 3). Defrost the required 4% PFA stock solution (each 96-well plate requires ~20 mL of solution). 6. Drug plate blocking (optional): Blocking the plate used to make up the test compounds with protein and nonionic detergent (bovine serum albumin and Tween-80) is required for the assessment of small molecule compounds on endocytosis pathways to reduce potential loss of the test compound due to non-specific plate binding, and ensuring accurate drug concentration in solution. Obtain and label 96-well polypropylene V-bottom microplate. Each of these wells (containing test compounds) will correlate with one drug condition in the fibronectin-coated glass-bottom plate (containing cells). Add 200 μL/well of blocking solution, and incubate at room temperature for at least 2 h. Prior to use, vigorously flick off the blocking solution and allow to air-dry. 7. Drug stock preparation (optional): Prepare drug stock(s) as 10 mM concentrations in 100% dimethyl sulfoxide (DMSO) if solubility permits. Label and create a vertical series of six concentrations (100, 30, 10, 3, 1, and 0.3 μM) for each

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Fig. 2 Schematic of 96-well drug plate representing the screening of four potential endocytosis inhibitors and the relevant dilutions required for each

drug, using 1.5 mL Eppendorf® Safe-Lock tubes in a microcentrifuge tube rack (see Notes 4 and 5). A template such as that illustrated is valuable to appropriately plan dilutions from each drug stock solution (Fig. 2). 8. Drug plate(s) preparation (optional): Pipette 2 μL/well of each drug stock (from lowest to highest concentration) to the drug plate(s) in accordance with the plate layout schematic, adding DMSO controls first (Fig. 2). 9. Drug incubation (optional): Sterile working conditions for the cells are no longer required. Following time elapse for serum starvation, add 198 μL/well of pre-warmed DMEM (without FBS) to the drug plate using a multichannel pipette (5–250 μL) with maximal output pressure to promote mixing (1:100 dilution to achieve final experimental drug concentration). Obtain MatriPlate™ cell plate from the incubator, and flick off media from cells. Using fresh pipette tips each time, transfer 180 μL/ well of the drug/DMEM solution from the drug plate(s) to the cell plate(s) following a pre-planned schematic plate layout. Place the plate(s) (with cells containing drug treatments) in the incubator (37  C, 5% CO2) for 30 min (see Note 6). 10. Ligand incubation: If not assessing the effect of small molecules on endocytic pathways, simply remove the media following serum starvation, and replace with 180 μL/well of fresh DMEM (to allow for fair comparison across experiments drug vs. non-drug experiments). The ligand to be assessed

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will determine which endocytosis pathway is to be studied: (1) transferrin or EGF (CME), (2) cholera toxin (CLIC/ GEEC), and (3) dextran 40 (macropinocytosis). Note that each ligand is not likely to be exclusively targeted to a single pathway but primarily uses the indicated path. Multiplexing at this stage can be established by applying different fluorescently labelled ligands, where the Alexa Fluor (AF) family of fluorescent dyes is particularly useful (AF 488-EGF and AF 594-transferrin). Do not overlap fluorophore excitation/emission channels across the chosen ligands and transfected proteins (if applicable). Dilute stock fluorescent ligand(s) solution in PBS to optimal dilution. Using a multichannel pipette (5–250 μL) with maximal output pressure, add 10 μL of diluted stock to each well, agitate the cell plate by gently tapping each side to disperse the ligand uniformly, and then incubate cells at 37  C. For example, transferrin uptake in U2OS cells can be achieved via addition of 10 μL of diluted AF 594-transferrin solution (4–5 μg/mL) for 8 min as this provides enough signal to assess the effect of potential endocytosis inhibitors (see Note 7). 11. Plate washing, fixation, and staining: Following ligand incubation, flick off the solution, and immediately place the 96-well cell plate on ice-cold water, ensuring cold water covers the entire base of the plate(s). Low temperature terminates endocytosis and subsequent endosomal trafficking processes and traps the endocytosed ligand inside the cells. Wash to remove extracellular ligand, fix, and stain the cells on the plate as per the following (see Note 8): (a) Add 200 μL/well of cold, low-pH acid wash solution, and incubate on ice for 10 min (removes extracellular membrane-bound transferrin ligand). Remove after time has elapsed. (b) Add 200 μL/well of cold PBS and incubate on ice for 5 min. Remove once time has elapsed, and place the plate at room temperature. (c) Add 200 μL/well of room temperature 4% PFA (pH 7.4) to fix the cells, and incubate at room temperature for 10 min. Remove PFA. (d) Make 50 mL of 200 μg/mL DAPI solution in PBS from concentrated DAPI stock (each 96-well plate requires ~20 mL of solution). Add 200 μL/well of diluted DAPI/PBS, and incubate at room temperature for 15 min to stain cell nuclei. Remove DAPI/PBS solution.

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(e) Add 200 μL/well of fresh PBS, wrap, and label individual plates in aluminum foil (to reduce photobleaching of fluorophores), and place in fridge until ready to proceed with imaging. Day 3: Plate image acquisition 12. IXM endocytosis imaging setup and acquisition: This protocol makes an assumption that a functional IXM system or alternative high-content imaging system is in operation with (1) access to DAPI and Texas Red excitation/emission capacities (with FITC for GFP transfection and transmitted light for morphology assessment), (2) a 20 Plan Fluor ELWD, and (3) accurate calibration. Prior to image acquisition, switch on IXM components (fans, lumencor, and sample plate temperature control), launch MetaMorph software, and equilibrate the endocytosis MatriPlate™ to room temperature (1 h prior to starting). Clean the glass bottom of the MatriPlate™ with ethanol and non-disintegrating absorbent tissues (e.g. Kimwipes) to remove any contaminant (dust/stains) capable of interfering with image acquisition. From the MetaMorph software home screen, select “Screening” followed by “Plate Acquisition Setup”. Given the high throughput, a 20 Plan Fluor ELWD magnification should be used (with 0.33  0.33 μm camera pixel binning). The plate dimensions are input into the IXM settings (ideal parameters are available from suppliers), with any deviation adjusted using “Autofocus” options. For the glass-bottom MatriPlate™, the following settings are applied: Number of Rows (8), Number of Columns (12), Well Shape (circle; assay established using circular area within square glass-bottom area), Well Diameter (7,000 μm), Column Spacing (9,000 μm), Plate Length (127.8 mm), Column Offset (14,380 μm), Row Spacing (9,000 μm), Plate Width (85.5 mm), Row Offset (11,240 μm), Well Depth (13,520 μm), and Plate Height (14.4 mm). Once the correct parameters have been entered, select “Sites to Visit”. Set “Site Acquisition Mode” to “Fixed Number of Sites” and adjust “Total Number of Sites” and their respective size (“Custom Field of View”) and spacing (μm) as desired. The example protocol here utilizes a centered tile configuration of 3  3 (columns  rows) with 300 μm even spacing to ensure large and distributed sample size. Given the static and fixed nature of the samples, under “Time lapse”, select a single time point (1). If the experiment does not require transfection (GFP-tagged protein), under “Acquisition Loop”, set “Number of Wavelengths” to 2 (DAPI, nucleus; Texas Red, transferrin) and select “Enable Laser-Based Focusing” within “Autofocus Options”. Next, under the

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“Autofocus” tab, employ the following settings to allow for bottom-of-well autofocusing of fluorophore signal (as physical variation between plates needs to be accounted for): Well to Well Autofocus (Focus on Well Bottom), Image-based Focusing (Algorithm, Standard; Binning 2), Initial Well for Finding Sample (First Well Acquired, 1), and Site Autofocus (All Sites). For wavelengths (W1/W2), select DAPI and Texas Red, and discover both the correct exposure (ms) using “Auto Expose” (DAPI, 250 ms; Texas Red, 4500 ms) and the appropriate “Laser with z-offset” using “Calculate Offset” option (using A1 DMSO control). Once focus and the correct exposure have been deciphered, return to “Autofocus” tab and “Configure Laser Autofocus Settings”. Choose to “Find Sample” as confirmation of correct focus and exposure. Once ready to initiate automated image acquisition, select “Summary” tab and “Save Settings” for future experiments as well as confirming all details are accurate (especially data storage capacity). Once confirmed, “Acquire Plate” and leave the IXM to run, returning only once the acquisition has been completed. For future experiments, a streamlined protocol can be followed. Select “Plate Acquisition Setup” and “Load Existing Settings File” to choose the saved settings. Provide a name and accurate description of the experimental setup, and select all the relevant wells to be analyzed. Shift the camera position to A1 (DMSO control; right-click well), click on “W1 DAPI” to change to the relevant laser and, within the “Autofocus” tab, opt to “Configure Laser Settings”. To ensure the plate is in proper focus with the correct offset, select to “Find Sample”, and a sample image within the well will be taken (revisit z-stack offset adjustment if not in focus). Repeat for “W2 Texas Red” to ensure both wavelengths are in focus; otherwise, analysis can be complicated. Click on “Summary” tab, and once the information has been reviewed, “Acquire Plate” to start automated image acquisition. If other plates are to be imaged, simply replace the MatriPlate™ and repeat. If desired, store the plate wrapped in aluminum foil at 4  C for future use. If moving on to image analysis, turn off the IXM components from Step 12 and relaunch the MetaMorph software. Day 4: Automated image and data analysis 13. Automated image analysis: Once quality, focused images have been acquired, analysis and quantification of endocytosis signal follow. An adaptation of the “Transfluor” module in-built within the MetaXpress software is used for quantification of the endocytosis signal. Launch MetaXpress software, select “Screening” and “Review Plate Data”. “Select Plate” recently acquired and choose to display “All Sites” of DAPI/Texas Red wavelengths to reveal a complete montage of images (Fig. 3).

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Fig. 3 Overview of 96-well plate image acquisition provides estimate of assay results. (a) Transferrin (AF 594; Texas Red) and nucleus (DAPI) signal of a typical 96-well plate format assessing four different compounds. The visual reduction in red indicates CME inhibition. Similarly, images that are mostly blue indicate CME inhibition, where cells remain attached to the plate. (b) Single-well snapshots of signals showing decrease of transferrin internalization upon increasing drug concentrations, while the cells remain attached to the plate

Under “Display” horizontal tab, deselect “color composite” if relevant, examine DAPI and Texas Red images for evidence of poor quality (e.g. lack of focus, drug toxicity/cell death indicated by unusual nuclear shapes or their complete absence; see Note 9), and annotate for later exclusion from analysis. Within “Run Analysis”, select “Transfluor” in “Analysis” drop-down menu followed by “Configure Settings”. Designate Texas Red to “Pits and Vesicles Image” and DAPI to “Nuclear Image”. Our protocol utilized the following

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Fig. 4 Automated image analysis module displaying nuclear mask (DAPI; green) and endocytosis signal (AF 594; Texas Red; pits/vesicles represented by white and red dots)

parameters (modify as necessary): transferrin pits (1–3 μm), transferrin vesicles (4–10 μm), nucleus (8–35 μm), and gray levels above background threshold (1000 for pits/vesicle and 500 for nuclear stain; test as necessary). Select “Test Run” to assess settings, and ensure automated analysis is quantifying observable signal, changing parameters (especially background threshold) as necessary (Fig. 4). “Save settings” for future analysis, choose to “Run on: All wells” and “Run Analysis”. 14. Data export: In the MetaXpress software, select “Screening” drop-down menu followed by “Review Plate Data”, and load the analyzed image plate. Within the “Measurements” tab, select the appropriate analysis (Transfluor: *Module Name*). To generate an inhibition curve, select “Open Log” and export “Transfluor: Nuclear Count” and “Cell: Vesicle Integrated Intensity (Transfluor)”. Various other useful analytical parameters can also be obtained from this analysis module. 15. Inhibition curve and analysis: The effect of endocytosis inhibitors is expressed as a percentage of the DMSO negative control at each respective concentration. If the desired target for a small molecule is suspected or known, secondary validation should be undertaken of the drug effect on that protein, given the large network of proteins required for endocytosis to occur (see Note 10). Divide the values of “Cell: Vesicle Integrated Intensity (Transfluor)” by their respective

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Fig. 5 Endocytosis inhibition curves displaying both active (dyngo®-4a, dyngo®6a, and dyngo®-7a) and inactive (dyngo®-1a) compound actions [19]

“Transfluor: Nuclear Count” values to obtain an average transferrin signal per cell. Assess DMSO controls (24 in total for 96-well plate), and exclude any obvious outliers (with justification following manual image review). Average the DMSO transferrin signal per cell values to obtain an overall DMSO control average. Proceed to divide the transferrin signal per cell from all treatment wells by the overall DMSO control average, and multiply by 100 to obtain a measurement of endocytosis as a percentage of the non-drug treatment DMSO wells. Using GraphPad Prism software, plot the average values from each of these percentages (three per condition) and SEM (standard error of mean) as a function of drug concentration (Fig. 5). Application of a nonlinear regression curve allows for the derivation of the associated half maximal inhibitory coefficient (IC50).

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Notes 1. Cell seeding density: Depending on the cell type, differences in size and morphology will affect how many cells are required to achieve confluency on any given surface. Endocytosis assay image acquisition and analysis require that there are no overlapping cells (80–90% confluency) so as to not complicate results and to ensure that the signal can be assigned to each individual cell with minimal influence from neighboring cells. Seeding of a serial dilution of a given cell type onto a coated plate followed by a quick endocytosis experiment (ligand incubation, fixation, and staining) can rapidly determine the correct

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seeding density for different cell types with variant growth rate characteristics. 2. Coating of glass-bottom 96-well plates: Although fibronectin is suitable for most cell lines, there are instances where different surfaces may be required (e.g. neurons typically prefer poly-Dlysine). If suspected that attachment is an issue, seed the same number of cells onto a variety of coating agents (e.g. gelatin, poly-D-lysine), perform the assay, and compare levels of cell attachment. 3. Serum starvation: Non-transfected and untreated (i.e. no siRNA or drug treatment) cells are normally capable of withstanding long periods of serum starvation without complication. However, if it is suspected that serum withdrawal may be an issue for cell function, the time frame can be shortened to at least 2 h. Plating of cells followed by a serum starvation time course and microscope investigation (looking for signs of cell rounding/death) will allow for suitable time frames to be established for each cell line or endocytosis pathway. Serum starvation is performed for endocytosis modes that are stimulated by ligands, such as CME, and the ligands added in a timed manner. However, the need for serum starvation must be determined for each endocytic mode. For example, FEME is wholly serum-dependent and stimulated by an increase in serum concentration [3]; hence, such cells are typically not serum starved. 4. Drug screening concentration range: The DMSO used for drug stock preparation should be fresh and of high quality. The solvent oxidizes upon storage, and it is recommended to use DMSO from sealed ampules for each experiment in order to reduce this technical problem. The dilution series shown here is loosely based on being linear when the data is represented on a log10 scale. Depending on the experiment, the drug dilution series can be adapted. For general purpose screening, the 0.3–100 μM range is suitable. However, for potent inhibitors, it is appropriate to utilize a shorter dilution range. Note that at high drug concentrations, there are risks to be aware of, notably, compound precipitation on the cells or cell lifting from the plate. 5. Drug selection for screening: The following two issues, inherent with the chemistry of some compound classes, make them unsuitable for this protocol: (1) fluorescence (ML141, a common cdc42 inhibitor, emits light in response to DAPI wavelength excitation, rendering nucleus-based automated cell counting impossible to perform; many experimental compounds are fluorescent across a broad spectrum) and (2) drug insolubility or aggregation (compounds may precipitate at

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higher concentrations or after dilution into the wells with the cells). If possible, a known positive control should be included with every batch of experiments to increase confidence that the assay itself is not the issue when such events unexpectedly arise. If inhibition of a particular pathway is desired, or if attempting to decipher through which pathway a given ligand is internalized, endocytosis inhibitors are available, each with a relative degree of specificity for certain pathways: (1) CME for dissection of dependence on clathrin (pitstop®-2) or dynamin (dyngo®-4a, dynole 34-2, chlorpromazine), (2) FEME (clathrin-independent (insensitive to pitstop®-2); no known specific inhibitors; however, it is blocked by dynamin modulators like dyngo®-4a and dynole 34-2 which also block CME), (3) caveolae-dependent endocytosis (filipin and β-cyclodextrin cholesterol modifiers), (4) CLIC/GEEC (dynamin and clathrin independent; no known specific inhibitors), (5) arf6associated endocytosis (arf6 peptide inhibitor), (6) rho-dependent IL-2 endocytosis (rhosin, a rho GTPase activation site inhibitor), (7) fluid phase endocytosis or macropinocytosis (amiloride, actin inhibitors), and (8) phagocytosis (actin inhibitors, rhoGEF inhibitors, dynamin inhibitors) [9– 12]. For multiplexing assays, inclusion of pitstop®-2 can reduce non-specific endocytosis through the CME pathway, while inclusion of dyngo®-4a can increase cargo specificity toward the CLIC/GEEC pathway. The use of actin inhibitors for multiplexing is not advisable due to its broad and sometimes inconsistent involvement in multiple pathways in a cell linedependent manner. 6. Duration of drug treatment: A compound incubation time of 30 min on the cells is generally suitable for most cell types, but if it is suspected that a different time frame is required (due to lack of effect of a known control or having significant issues with cell lifting), perform a time course of drug treatments to establish optimal conditions. Active compounds can be checked later for optimal drug exposure duration. 7. Conditions for ligand incubation: Depending on the cell type and the chosen ligand in question, incubation times will need to vary to accommodate differential ligand internalization (fluorescent signal). Ligands with poor internalization by a cell type will require a longer incubation time to confidently note a compound effect. Both a time course and an assessment of different fluorescence ligand concentrations will allow for establishment of optimal conditions. Such preliminary experiments also need to consider the trafficking of the internalized ligand, some of which may be rapidly exocytosed (e.g. transferrin) or accumulate in a perinuclear location. If multiplexing ligands, simply add the ligand with the

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Fig. 6 Manual image review reveals technical or cytotoxicity issues. (a) Out-offocus-well snapshot (blurry). (b) Compound fluorescence (entire cell content is fluorescent). (c) Cell toxicity (smaller nuclei). (d) Compound aggregation (speckled appearance across the plate, not confined to cells). Each image is displayed at the same magnification

pre-determined longest incubation time, and wait until time elapses to add the other ligand and start the shorter incubation time (i.e. both will conclude at the same time). On the other hand, if the goal is to determine whether two ligands might be co-endocytosed, they should be added simultaneously, and the internalization time should be shortened (e.g. 2–4 min). For example, transferrin and EGF (low concentrations) are co-endocytosed by CME but exhibit divergent trafficking pathways within minutes of entry. 8. Cell plate washing: Low-pH acid wash typically releases ligands such as transferrin from their receptor, reducing the detection of extracellular receptors to near background. The low-pH washing steps listed are widely used and should be well tolerated by most cell types, but if it is noted that a cell type is sensitive, take the appropriate precautions; for example, the acid wash pH could potentially be increased in pH and yet still have a similar effect of removing membrane-bound ligands. GFP denatures and loses fluorescence under pH 6.0 so take care upon transfection.

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9. Manual image review: To ensure confidence in the automated analysis results, a manual review of all the input images should be undertaken (Fig. 6). There exists several points of observation: (1) image focus (blurry snapshots can interrupt automated quantification), (2) drug fluorescence (if the test drug is intrinsically fluorescent, it can interfere with automated analysis by overlapping with the DAPI mask or the Texas Red marker of vesicles and endosomes), (3) nuclear DAPI stain (a lack of any DAPI staining of cells can indicate cell lifting by the drug, while lack of uniformity in nucleus size and shape is an indicator of drug toxicity; if necessary, a colorimetric MTT dye assay can be utilized to quantify cell viability), and (4) drug aggregation (capable of trapping endocytic cargo and complicating analysis). 10. Secondary validation of endocytosis target: The endocytosis high-content screening analysis described here is a functional or phenotypic screen. It reports endocytosis or trafficking actions of potential drugs but does little to indicate the target of the drug. If possible and desired, given the multifactorial nature of endocytosis, validation of the small molecule target can be achieved through other methods (i.e. in vitro biochemistry). Assays have been reported in the literature that can specifically assess drug action upon dynamin GTPase activity or clathrin protein-protein interactions [13, 14]. Endocytosis by CME is highly sensitive to inhibiting the function of either of these two proteins. Either or both may also be off-target action of compounds presumed or known to target other endocytic proteins (e.g. phenothiazines such as chlorpromazine are reported to bind AP2 but block dynamin GTPase activity [15]). A significantly lower endocytosis inhibition coefficient (IC50) can be suggestive of multiple endocytic targets of a given small molecule. A further possible avenue of validation involves taking advantage of the spontaneous strainpromoted alkyne-azide cycloaddition reaction in a cellular context. This involves four central steps: (1) synthesis of an azide analogue of the small molecule of interest, (2) azide drug treatment of cells overexpressing the suspected protein target (drug-protein interaction), (3) incubation of cells with DBCOconjugated fluorophore (resulting in azide-alkyne covalent attachment), and (4) fixation, staining, and confocal imaging should see co-localization between protein and fluorophore. Another approach involves a cellular thermal shift assay, a mass spectrometry-based and label-free approach of determining drug-binding events in cells and tissues [16]. Overall, it can be challenging to determine which protein is the authentic target of a drug that inhibits endocytosis in a phenotypic screen.

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Table 1 Summary of high-content drug screening protocol advantages and disadvantages Advantages

Disadvantages

Information-rich

Requires access to high-content imaging microscope

High throughput

Unsuitable for fluorescent test compounds

Straightforward and versatile protocol

Molecular target of test compound can remain unclear

Multiplexable to numerous ligands or endocytic pathways

Difficult to discriminate impact of test compound on cell lifting vs. cytotoxicity

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Conclusion Identification of pathway-specific endocytosis modulators has broad clinical utility, with potential applications as diverse as epilepsy and viral particle entry. The broad-spectrum, multiplexable, and high-content screening assay illustrated here is capable of identifying potential small molecule candidates for endocytosis inhibition. This protocol exhibits several key advantages and disadvantages, with one major limitation being its inability to unequivocally identify the target of such small molecules, which remains a challenge for all drug discovery (Table 1) [17]. Although this protocol focuses on transferrin-mediated CME using the unmodified U2OS cell line, the procedure is highly adaptable to a multitude of ligands (single or multiplexed), cell lines (native, transfected, or genetically engineered to lack or replace a single protein), and/or treatment conditions, depending on the experimental question (with a range of troubleshooting and general notes provided to accommodate). Multiplexing is valuable in determining the cross action of a compound on another pathway, thus reporting on its specificity. For example, dynamin inhibitors block CME (of transferrin and low concentrations of EGF), as well as FEME (of high concentrations of EGF, and many other ligands [3, 18]), while pitstop®-2 inhibits only CME. Judicious application of pitstop®-2 (10–20 μM) to the assay described here to reduce CME can be valuable in reporting the specificity of other nonclathrin-mediated endocytic pathways. Dynamin inhibitors can be utilized in a similar manner. Since no inhibitor is completely specific to a single pathway, this approach can increase the specificity of detection of non-dynamin, non-clathrin dependent endocytic routes in multiplexed functional assays. In regards to the early compound research and development phase, this method provides three primary uses: (1) identification of novel scaffolds capable of endocytosis inhibition, (2) structureactivity relationship assessment of analogues in pursuit of developing more potent compounds, and (3) in an instance where

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endocytosis inhibition as an off-target is not desired, novel compounds can be assessed and cleared. For fundamental research purposes, this protocol can also be extended to assess internalization pathways of ligands as well as establish the importance of certain proteins/mutants in a given pathway (via transfection, CRISPR-Cas9 knockout, or siRNA knockdown). This methodology provides a powerful platform with which to answer many experimental questions in both a research and industrial context.

Acknowledgments We are grateful for financial support from the National Health and Medical Research Council Australia (GNT1069493, GNT1052494, GNT1047070, GNT 1105666, GNT1137064, and GNT1162515), Australian Research Council (DP180101781), Children’s Medical Research Institute (CMRI), and the University of Sydney and for equipment from the Australian Cancer Research Foundation, the Ramaciotti Foundation, and the Cancer Institute NSW. Endocytosis pathways (Fig. 1) were created with BioRender.com. References 1. McMahon HT, Boucrot E (2011) Molecular mechanism and physiological functions of clathrin-mediated endocytosis. Nat Rev Mol Cell Biol 12:517–533 2. Doherty GJ, McMahon HT (2009) Mechanisms of endocytosis. AnnuRevBiochem 78:857–902 3. Boucrot E, Ferreira AP, Almeida-Souza L, Debard S, Vallis Y, Howard G, Bertot L, Sauvonnet N, McMahon HT (2015) Endophilin marks and controls a clathrin-independent endocytic pathway. Nature 517:460–465 4. Harper CB, Popoff MR, McCluskey A, Robinson PJ, Meunier FA (2013) Targeting membrane trafficking in infection prophylaxis: dynamin inhibitors. Trends Cell Biol 23:90–101 5. Li YY, Chen XN, Fan XX, Zhang YJ, Gu J, Fu XW, Wang ZH, Wang XF, Xiao Z (2015) Upregulated dynamin 1 in an acute seizure model and in epileptic patients. Synapse 69:67–77 6. Luwor RB, Morokoff AP, Amiridis S, D’Abaco G, Paradiso L, Stylli SS, Nguyen HPT, Tarleton M, Young KA, O’Brien TJ, Robinson PJ, Chircop M, McCluskey A, Jones NC (2019) Targeting glioma stem cells by functional inhibition of dynamin 2: a novel

treatment strategy for glioblastoma. Cancer Investig 25:1–12 7. Harper CB, Martin S, Nguyen TH, Daniels SJ, Lavidis NA, Popoff MR, Hadzic G, Mariana A, Chau N, McCluskey A, Robinson PJ, Meunier FA (2011) Dynamin inhibition blocks botulinum neurotoxin type-A endocytosis in neurons and delays botulism. J Biol Chem 286:35966–35976 8. Jo U, Park KH, Whang YM, Sung JS, Won NH, Park JK, Kim YH (2014) EGFR endocytosis is a novel therapeutic target in lung cancer with wild-type EGFR. Oncotarget 5:1265–1278 9. Ivanov AI (2008) Pharmacological inhibition of endocytic pathways: is it specific enough to be useful? Methods Mol Biol 440:15–33 10. Otsuka A, Abe T, Watanabe M, Yagisawa H, Takei K, Yamada H (2009) Dynamin 2 is required for actin assembly in phagocytosis in Sertoli cells. Biochem Biophys Res Commun 378:478–482 11. Shang X, Marchioni F, Sipes N, Evelyn CR, Jerabek-Willemsen M, Duhr S, Seibel W, Wortman M, Zheng Y (2012) Rational design of small molecule inhibitors targeting RhoA subfamily Rho GTPases. Chem Biol 19:699–710

High-Content Drug Screening of Endocytosis Pathways 12. Davis CT, Zhu W, Gibson CC, BowmanKirigin JA, Sorensen L, Ling J, Sun H, Navankasattusas S, Li DY (2014) ARF6 inhibition stabilizes the vasculature and enhances survival during endotoxic shock. J Immunol 192:6045–6052 13. Quan A, Robinson PJ (2005) Rapid purification of native dynamin I and colorimetric GTPase assay. Methods Enzymol 404 (Ch 49):556–569 14. von Kleist L, Stahlschmidt W, Bulut H, Gromova K, Puchkov D, Robertson MJ, MacGregor KA, Tomilin N, Pechstein A, Chau N, Chircop M, Sakoff J, von Kreis J, Saenger W, Krausslich HG, Shupliakov O, Robinson PJ, McCluskey A, Haucke V (2011) Role of the clathrin terminal domain in regulating coated pit dynamics revealed by small molecule inhibition. Cell 146:471–484 15. Daniel JA, Chau N, Abdel-Hamid MK, Hu L, von Kleist L, Whiting A, Krishnan S, Maamary P, Joseph SR, Simpson F, Haucke V, McCluskey A, Robinson PJ (2015) Phenothiazine-derived antipsychotic drugs inhibit dynamin and clathrin-mediated endocytosis. Traffic 15:635–654 16. Molina DM, Nordlund P (2016) The cellular thermal shift assay: a novel biophysical assay for in situ drug target engagement and

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mechanistic biomarker studies. Annu Rev Pharmacol Toxicol 56:141–161 17. Lin A, Giuliano CJ, Palladino A, John KM, Abramowicz C, Yuan ML, Sausville EL, Lukow DA, Liu L, Chait AR, Galluzzo ZC, Tucker C, Sheltzer JM (2019) Off-target toxicity is a common mechanism of action of cancer drugs undergoing clinical trials. Sci Transl Med 11:eaaw8412 18. Renard HF, Simunovic M, Lemiere J, Boucrot E, Garcia-Castillo MD, Arumugam S, Chambon V, Lamaze C, Wunder C, Kenworthy AK, Schmidt AA, McMahon HT, Sykes C, Bassereau P, Johannes L (2015) Endophilin-A2 functions in membrane scission in clathrin-independent endocytosis. Nature 517:493–496 19. McCluskey A, Daniel JA, Hadzic G, Chau N, Clayton EL, Mariana A, Whiting A, Gorgani N, Lloyd J, Quan A, Moshkanbaryans L, Krishnan S, Perera S, Chircop M, von Kleist L, McGeachie AB, Howes MT, Parton RG, Campbell M, Sakoff JA, Wang X, Sun JY, Robertson MJ, Deane FM, Nguyen TH, Meunier FA, Cousin MA, Robinson PJ (2013) Building a better dynasore: the dyngo compounds potently inhibit dynamin and endocytosis. Traffic 14:1272–1289

Chapter 6 Methods for Monitoring Endocytosis in Astrocytes Maja Potokar, Jernej Jorgacˇevski, and Robert Zorec Abstract Endocytosis is a vesicle-based mechanism by which eukaryotic cells internalize extracellular material. There are several types of this universal mechanism linked to different types of endocytosed cargo, including pathogens; therefore, several approaches can be applied. Here, we describe techniques that are applicable to study the internalization of flaviviruses; dextrans; transporters, such as, glutamate transporter vGlut1; and peptidergic signaling molecules, including atrial natriuretic peptide into astrocytes, the most heterogeneous neuroglial cells, which play a key homeostatic role in the central nervous system. Key words Endocytosis, Recycling vesicles, Astrocyte, Flavivirus, vGlut1, ANP, Gliotransmitter, Dextran

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Introduction Endocytosis is the general term for internalization of fluids, solutes, macromolecules, plasma membrane components, and different particles, including extracellular pathogens by invagination of the plasma membrane and the formation of cytoplasmic vesicles and vacuoles through membrane fission [1]. In this way, metazoan cells endocytose nutrients, receptor-ligand complexes, fluids, solutes, lipids, membrane proteins, extracellular matrix components, cell debris, bacteria, and viruses, that, by passing through endosomal vesicles, affect regulation and fine-tune numerous pathways in the cell [1]. Astrocytes are the neural cell type crucially involved in homeostasis of the central nervous system, control of synaptogenesis, synaptic connectivity, integration and synchronization of neuronal networks, and the maintenance of the integrity of the blood-brain barrier and are part of the glymphatic system that contributes to the regulation of waste removal from the extracellular space [2, 3]. In addition, astrocytes play an important phagocytic role by engulfing synapses, apoptotic cells, cell debris, and released toxic proteins and therefore have a critical role in the development of neurodegenerative diseases [4, 5]. Trafficking of

Florence Niedergang, Nicolas Vitale and Ste´phane Gasman (eds.), Exocytosis and Endocytosis: Methods and Protocols, Methods in Molecular Biology, vol. 2233, https://doi.org/10.1007/978-1-0716-1044-2_6, © Springer Science+Business Media, LLC, part of Springer Nature 2021

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endocytotic and recycling vesicles in astrocytes has been monitored in several experimental settings, and various mobility parameters have been described [6–9]. Thus, we focus here on four labelling techniques applicable for observing endocytosis of flaviviruses, dextrans, vesicular glutamate transporter 1 (vGlut1), and atrial natriuretic peptide (ANP) [6, 7, 10–12].

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Materials All reagents should be of analytical grade. In all labelling steps, it is mandatory to use solutions prepared with MilliQ water. 1. Phosphate-Buffered Saline (PBS): 0.01 M phosphate buffer, 0.0027 M potassium chloride, and 0.137 M sodium chloride (pH 7.4) at 25  C (see Note 1). 2. Artificial CFS (aCSF): 124 mM NaCl, 3.1 mM KCl, 1.25 mM NaH2PO4·H2O, 26 mM NaHCO3, 10 mM D-glucose, 1 mM MgCl2, and 2 mM CaCl2 at pH 7. Aerate the solution with 95% O2/5% CO2 (see Note 2). 3. Solution for diluting antibodies: 3% bovine serum albumin (BSA) in PBS (see Note 3). 4. Extracellular solution: 130 mM NaCl, 2 mM CaCl2, 5 mM KCl, 1 mM MgCl2, 10 mM HEPES, and 10 mM D-glucose (pH 7.2, 300 mOsm/L). 5. HEPES Buffer: 50 mM HEPES and 145 mM NaCl (pH 7.4).

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Methods Labelling and washing steps should be carried out at room temperature (RT), unless otherwise specified.

3.1 Endocytosis of Flaviviruses

Laboratory workers may be exposed to a risk of viral infections that could result in severe illnesses for which there may be unproven treatments or none at all. Research facilities and individual researchers contemplating work with pathogens should be mindful of the occupational health issues, and all experiments requiring manipulation of viruses should be undertaken in facilities with an appropriate biosafety level. 1. Grow flavivirus for 4–7 days on Vero E6 cells, and collect virus supernatant into 1 mL microcentrifuge tubes. 2. Centrifuge at 3000  g (10 min, 4  C) (see Note 4). 3. Transfer the supernatant into fresh 1 mL microcentrifuge tubes. 4. Centrifuge at 20,000  g (5 min, 4  C) (see Note 4).

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Fig. 1 DiD-TBEV (tick-borne encephalitis virus) particles in astrocytes are localized in endosomes and lysosomes. (ai, ii). An astrocyte with DiD-labelled TBEV particles (TBEV) incubated at 37  C for 4 h and immunolabelled with antibodies against early endosomal antigen (EEA1, 1:300), a marker for early endosomes. Overlays represent overlapped DiD-TBEV and EEA1 fluorescent signals, indicating the association between DiD-TBEV and endosomes. Scale bars: 5 μm. (bi, ii) An astrocyte with DiD-labelled TBEV vesicles (TBEV) incubated at 37  C for 4 h and immunolabelled with antibodies against lysosomal-associated membrane protein 1 (LAMP1; 1:300), a marker for late endosomes/lysosomes. Overlays represent overlapped DiD-TBEV and LAMP1 fluorescent signals, indicating the association between DiD-TBEV and late endosomes/ lysosomes. Scale bars: 5 μm. (aiii, biii). Prolonged incubation significantly increased the average number of DiD-TBEV-labelled vesicles per cell. *P < 0.05; ***P < 0.001. (Modified from Ref. 12 with permission)

5. Transfer the supernatant into fresh 1 mL microcentrifuge tubes, and add fluorescent lipophilic dye at an optimal, empirically determined concentration (see Note 5). Incubate for 2 h at 37  C in a heated shaker. 6. Remove unbound dye by buffer exchange into HEPES buffer. Use disposable columns for rapid and efficient purification of DNA and oligonucleotides (>10-mer) prepacked with Sephadex G-25 DNA Grade. 7. Aliquot the labelled virus and proceed with infection of astrocytes. Store remaining aliquots at 80  C.

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8. Dilute labelled virus at the desired multiplicity of infection (see Note 6) in growth medium, and apply the mixture to astrocytes. The mixture should cover plated cells (see Note 7) at 4  C for 10 min to synchronize virus infection and then incubate at 37  C for 1 h. 9. Wash cells 2 in PBS. 10. Apply fresh growth medium, and incubate at 37  C for the preferred time after infection. 11. Record endocytosed virus in astrocytes in a tightly sealed recording chamber (see Note 8, Fig. 1). 3.2 Endocytosis of Dextrans in Astrocytes

1. Briefly rinse cells in PBS. 2. Dilute dextrans in cell growth medium (see Note 9). 3. Apply diluted dextrans to astrocytes and incubate at 37  C for 16 h. 4. Rinse with Dulbecco’s Modified Eagle Medium. 5. Replace medium with fresh growth medium and incubate for 3 h at 37  C. 6. Record endocytosed dextrans in extracellular solution. Alternatively, you can fix samples and label them by immunocytochemical staining (see Note 10). Travelling through the endocytotic pathway, most endocytosed dextrans localize in late endosomes/lysosomes after 19 h of incubation (Fig. 2).

3.3 Labelling of Endocytotic Vesicles in Live Astrocytes with Antibodies

1. Briefly rinse cells in PBS. 2. Incubate cells in 3% BSA in PBS (2 min, RT) (see Note 11). 3. Label cells with primary antibodies diluted in 3% BSA in PBS (10 min, RT) (see Note 12). Incubate control samples only in 3% BSA in PBS (see Note 13). 4. Rinse cells in PBS (3, 2 min, RT). 5. Label cells with secondary antibodies diluted in 3% BSA in PBS (20 min, RT). Rinse cells in PBS (3, 2 min, RT). From here, you can follow steps 6 or 7. 6. Transfer samples to the recording chamber, apply extracellular solution, and observe endocytotic vesicles in real time (see Fig. 3 for an example of labelled endocytotic vesicles in live cultured astrocytes). 7. Fix cells in 4% formaldehyde (15 min, RT), and follow the immunostaining protocol (see Note 14).

3.4 Labelling of Endocytotic Vesicles in Astrocytes in Acute Tissue Slices with Antibodies

1. Transfer tissue slices into the labelling chamber with aCSF (see Note 15). 2. Incubate tissue slices in 3% BSA in aCSF (2 min, RT) (see Note 11).

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Fig. 2 Dextrans are localized in late endosomes/lysosomes in astrocytes after 19 h of application of dextrans. Most dextran puncta are colocalized in late endosomes/lysosomes (overlay), as depicted in the panels displaying astrocytes with endocytosed Alexa Fluor 546 dextrans, immunolabelled with antibodies against lysosomal-associated membrane protein 1 (LAMP1). Scale bars: 10 μm (2 μm in insets). Arrowhead points to typical structures expressing both signals. Note that the green signal corresponding to the membrane-bound LAMP-1 signal encircles the luminal signal of red dextran. (Modified from Ref. 10 with permission)

3. Label tissue slices with primary antibodies diluted in 3% aCSF in PBS (10 min, RT) (see Note 12). Incubate control samples only in 3% BSA in aCSF (see Note 13). 4. Rinse tissue slices in aCSF (3, 2 min, RT). 5. Label tissue slices with secondary antibodies diluted in 3% BSA in aCSF (20 min, RT). Rinse slices in aCSF (3, 2 min, RT). 6. Transfer tissue slices into the recording chamber, apply aCSF and observe endocytotic vesicles in real time (see Fig. 3 for an example of labelled endocytotic vesicles in live tissue slices).

4

Notes 1. For monitoring endocytosis in cell cultures, dilute reagents and wash samples in PBS. 2. For monitoring endocytosis in tissue slices, dilute reagents and wash samples in aCSF. 3. You can store aliquots of 3% BSA in PBS at thaw and warm them to RT before use.

20  C and then

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Fig. 3 Endocytosis of antibody-labelled vesicles/granules in live cultured astrocytes and brain slices. The procedure for vesicle/granule labelling with antibodies is presented in (ai–iii). Primary antibodies from the extracellular solution bind to extracellularly exposed vesicle/granule proteins of endocytosed/recycling vesicles (ai). Secondary antibodies bind to primary antibodies in one of the following vesicle/granule cycles (aii). Vesicles/granules labelled with both antibodies (aiii) can be imaged. Examples of vesicles/granules successfully labelled with anti-ANP primary antibodies and anti-rabbit Alexa Fluor secondary antibodies are depicted in (b). (bi) Labelling in cultured astrocytes (scale bar, 2.5 μm). (bii and iii) Labelling of ANP granules (bii) and vGlut1 granules (biii) in tissue slices (scale bar, 2.5 μm). (Modified from Ref. 13 with permission)

4. Cells and cell debris are removed from the supernatant with this step. 5. In our studies [11, 12], we labelled flaviviruses with 50 μM fluorescent lipophilic Vybrant DiD labelling solution that was freshly prepared from a 1 mM stock concentration. 6. The multiplicity of infection or MOI represents the ratio between the number of virus particles and the number of cells. A value of MOI 1 implies that on an average, a single virus particle infects a single cell. In example, assuming that if we apply 1 mL of 1  107/mL viral particles to 1  107 cells, we infect cells at an MOI 1. You can infect cells at a different MOI, adjusted to your experiment. 7. Astrocytes should be plated in recording dishes that can be tightly sealed for observation under the microscope. Ideally, plate astrocytes 24–48 h before infection. 8. To determine the localization of endocytosed virus particles, we immunolabelled cells by early and late endocytotic markers. Immunolabelling was performed after internalization of DiD virus particles. We fixed cells in 4% formaldehyde, washed them in PBS (4, 3 min, RT), incubated in blocking solution (goat serum, 1 h at 37  C), removed goat serum and labelled with secondary antibodies (45 min, 37  C), washed in PBS (3,

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3 min, RT), and mounted on glass slides in solution, which prevents fading of fluorophores [12]. The average number of DiD labelled virus particles per cell increased after longer time postinfection. The rate of virus endocytosis depends on the virus species and strain (Fig. 1) [11, 12]. 9. Previously prepare a fresh stock concentration of dextrans in 0.01 M PBS. In our study, we labelled astrocytes with Alexa Fluor 546 dextran conjugate at a final concentration of 0.1 μg/ mL [10]. 10. Pay extra attention to the fixation step. Fix samples with 2% formaldehyde for 5 min at RT. In case even this fixation proves to be to rigorous, the authors suggest further reducing the incubation time. 11. Incubating cells in 3% BSA in PBS (or aCSF) will reduce nonspecific labelling with antibodies. 12. Proper dilutions of antibodies should be determined on the basis of recommended dilutions for immunocytochemical staining provided by the antibody manufacturers. 13. Control samples are used to verify the absence of antibody labelling at the properly determined concentration of antibodies. 14. To further determine the localization of endocytotic vesicles (i.e., endocytotic markers, cytoskeleton markers), you can double label them with additional antibodies. After fixation in 4% formaldehyde, wash cells in PBS (4, 3 min, RT), incubate in blocking solution (i.e., goat serum, 1 h at 37  C), remove the goat serum and label with second primary antibodies (2 h at 37  C or overnight at 4  C) followed by secondary antibodies (45 min, 37  C), wash in PBS (3, 3 min, RT), and mount on glass slides in solution, which prevents fading of fluorophores. Store immunolabelled samples at 4  C. 15. Keep tissue slices in aerated solutions during the labelling and washing steps and during recording.

Acknowledgments This work was supported by the Research Agency of Slovenia (grant numbers P3 0310, J3 9266, and J3 7605), BI-JP/18-20-002, and EuroCellNet COST Action (CA15214).

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References 1. Huotari J, Helenius A (2011) Endosome maturation. EMBO J 30(17):3481–3500. https:// doi.org/10.1038/emboj.2011.286 2. Zorec R, Parpura V, Verkhratsky A (2018) Astroglial vesicular network: evolutionary trends, physiology and pathophysiology. Acta Physiol (Oxf) 222(2):e12915. https://doi. org/10.1111/apha.12915 3. Verkhratsky A, Nedergaard M (2016) The homeostatic astroglia emerges from evolutionary specialization of neural cells. Philos Trans R Soc Lond Ser B Biol Sci 371(1700):20150428. https://doi.org/10.1098/rstb.2015.0428 4. Tremblay ME, Cookson MR, Civiero L (2019) Glial phagocytic clearance in Parkinson’s disease. Mol Neurodegener 14(1):16. https:// doi.org/10.1186/s13024-019-0314-8 5. Morizawa YM, Hirayama Y, Ohno N, Shibata S, Shigetomi E, Sui Y, Nabekura J, Sato K, Okajima F, Takebayashi H, Okano H, Koizumi S (2017) Reactive astrocytes function as phagocytes after brain ischemia via ABCA1mediated pathway. Nat Commun 8(1):28. https://doi.org/10.1038/s41467-01700037-1 6. Stenovec M, Kreft M, Grilc S, Potokar M, Kreft M, Pangrsˇicˇ T, Zorec R (2007) Ca(2+)dependent mobility of vesicles capturing antiVGLUT1 antibodies. Exp Cell Res 13 (18):3809–3818. https://doi.org/10.1016/j. yexcr.2007.08.020 7. Potokar M, Stenovec M, Kreft M, Kreft M, Zorec R (2008) Stimulation inhibits the mobility of recycling peptidergic vesicles in astrocytes. Glia 56(2):135–144

8. Potokar M, Stenovec M, Kreft M, Gabrijel M, Zorec R (2011) Physiopathologic dynamics of vesicle traffic in astrocytes. Histol Histopathol 26(2):277–284 9. Potokar M, Vardjan N, Stenovec M, Gabrijel M, Trkov S, Jorgacˇevski J, Kreft M, Zorec R (2013) Astrocytic vesicle mobility in health and disease. Int J Mol Sci 14 (6):11238–11258. https://doi.org/10.3390/ ijms140611238 10. Vardjan N, Gabrijel M, Potokar M, Svajger U, Kreft M, Jeras M, de Pablo Y, Faiz M, Pekny M, Zorec R (2012) IFN-γ-induced increase in the mobility of MHC class II compartments in astrocytes depends on intermediate filaments. J Neuroinflammation 9:144. https://doi.org/ 10.1186/1742-2094-9-144 11. Jorgacˇevski J, Korva M, Potokar M, Lisjak M, ˇ upanc T, Zorec R (2019) ZIKV strains Avsˇicˇ-Z differentially affect survival of human fetal astrocytes versus neurons and traffic of ZIKVladen endocytotic compartments. Sci Rep 9 (1):8069. https://doi.org/10.1038/s41598019-44559-8 12. Potokar M, Korva M, Jorgacˇevski J, Avsˇiˇ upanc T, Zorec R (2014) Tick-borne cˇ-Z encephalitis virus infects rat astrocytes but does not affect their viability. PLoS One 9(1): e86219. https://doi.org/10.1371/journal. pone.0086219 13. Potokar M, Kreft M, Lee S, Takano H, Haydon P, Zorec R (2009) Trafficking of astrocytic vesicles in hippocampal slices. Biochem Biophys Res Commun 390(4):1192–1196. https://doi.org/10.1016/j.bbrc.2009.10. 119

Chapter 7 Monitoring Activity-Dependent Bulk Endocytosis in Primary Neuronal Culture Using Large Fluorescent Dextrans Michael A. Cousin and Karen J. Smillie Abstract The efficient recycling of synaptic vesicles (SVs) during neuronal activity is central for sustaining brain function. During intense neuronal activity, the dominant mechanism of SV retrieval is activity-dependent bulk endocytosis (ADBE). Here, we describe a method to monitor ADBE in isolation from other SV endocytosis modes, via the uptake of large fluorescent fluid-phase markers in primary neuronal culture. Furthermore, we outline how to monitor ADBE using this approach across a field of neurons or in individual neurons. Key words Dextran, Endocytosis, Vesicle, Neuron, Presynapse

1

Introduction The localized endocytosis of synaptic vesicles (SVs) at the presynapse is essential to sustain neurotransmission in mammalian brain. Three modes of endocytosis maintain SV supply, which are triggered by specific patterns of neuronal activity. During very mild neuronal activity, ultrafast endocytosis (UFE) is the dominant SV endocytosis mode. UFE forms small endosomes directly from the plasma membrane, before a second SV generation step [1, 2]. This pathway rapidly saturates during action potential trains, meaning it will likely provide a minor contribution to SV supply during physiological patterns of activity [3]. Action potential trains also trigger clathrin-mediated endocytosis (CME) [4]; however, this mode also saturates during periods of high neuronal activity [5]. During high neuronal activity, a different endocytosis mode is triggered, called activity-dependent bulk endocytosis (ADBE). ADBE is a two-step process, with large invaginations forming bulk endosomes directly from the plasma membrane [5], with SVs budding from these endosomes thereafter [6].

Florence Niedergang, Nicolas Vitale and Ste´phane Gasman (eds.), Exocytosis and Endocytosis: Methods and Protocols, Methods in Molecular Biology, vol. 2233, https://doi.org/10.1007/978-1-0716-1044-2_7, © Springer Science+Business Media, LLC, part of Springer Nature 2021

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Fig. 1 Overview of the tetramethylrhodamine (TMR)-dextran uptake assay to monitor activity-dependent bulk endocytosis (ADBE). Graphic shows 40 kDa TMR-dextran molecules in red which, following neuronal stimulation, are not accumulated into single retrieving synaptic vesicles but can be accumulated into larger endosomes formed by ADBE. This results in labelled endosomes which can then be detected by fluorescent imaging, selectively identifying nerve terminals which have undergone ADBE

The physiological role of ADBE in neuronal function is still undetermined, mainly because molecules specific to this process are only now starting to be identified [7]. Central to defining the role of key molecules in ADBE are assays that only report the triggering of this mode rather than either UFE or CME. Commonly used approaches to monitor endocytosis with either fluorescent dyes such as FM1-43 [8] or genetically encoded reporters such as synapto-pHluorins [9] are not appropriate, since they report the retrieval of membrane or proteins that are retrieved by all three endocytosis modes. One assay that monitors ADBE specifically is the internalization of large fluid-phase markers such as dextrans. High molecular weight versions of these molecules (40 kDa and 70 kDa) have difficulty entering single SVs formed via CME, meaning they almost exclusively report ADBE (Fig. 1) [10]. The extent of dextran labelling of UFE will be minimal, since this endocytosis mode provides a very minor contribution to SV retrieval during high neuronal activity [3]. Therefore, high molecular weight dextran molecules provide a simple optical estimation of the number of nerve terminals performing ADBE. This assay can be used for individual neurons or populations of neurons [7, 11–13]. A population-based approach is usually used when all neurons in primary culture are treated in a similar manner, for example, after application of a drug. It is also used to compare between cultures derived from animals with different genotypes or after viral transduction. The single neuron approach is typically used after transient transfection with either shRNA or overexpression plasmids, since the low transfection efficiency of this delivery method negates a population-based response.

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Materials Equipment

1. Sealed imaging chamber with embedded platinum wires such as a Warner imaging chamber (Warner RC-21BRFS). 2. Electrical field stimulator (see Note 1). 3. Perfusion system, for example, a gravity flow system with peristaltic pump. 4. Inverted epifluorescence microscope with standard rhodamine filters (plus appropriate cyan fluorescent protein (CFP)/green fluorescent protein (GFP) filters if required to identify individual transfected neurons) and a high numerical aperture (NA)
20 or
40 objective (see Note 2). A cooled chargedcoupled device digital camera connected to the microscope plus the associated operating software will also be required. 5. Primary cerebellar granule neurons (or other primary neuronal cultures) cultured on coverslips compatible with the imaging chamber (see Note 3). 6. Fiji (https://imagej.net/Fiji) and Microsoft Excel or other similar software to analyze the gathered images.

2.2 Solutions and Reagents

1. Imaging buffer for cerebellar granule neurons: 170 mM NaCl, 3.5 mM KCl, 0.4 mM KH2PO4, 20 mM TES (N-tris (hydroxyl-methyl)-methyl-2-aminoethane-sulfonic acid), 5 mM NaHCO3, 5 mM glucose, 1.2 mM MgCl2, and 1.3 mM CaCl2, pH 7.4 (see Note 4). (If using high-potassium solution (see Notes 1 and 11) instead of electrical stimulation, high-potassium imaging buffer is as above but with the substitution of 50 mM KCl for 50 mM NaCl.) 2. Imaging buffer for cortical/hippocampal neurons (if required): 136 mM NaCl, 2.5 mM KCl, 2 mM CaCl2, 1.3 mM MgCl2, 10 mM glucose, and 10 mM HEPES (2-[4-(2-hydrosxyethyl) piperazine-1-yl]ethanesulfonic acid), pH 7.4, supplemented with 10 μM CNQX (6-cyano-7-nitroquinoxaline-2,3-dione) and 50 μM APV (DL-2-amino-5-phosphopentanoic acid). 3. Dextran solution: 2 mM of 40 kDa TMR (tetramethylrhodamine)-dextran: in water (40
stock) stored at 20  C until use (see Note 5). 4. Vacuum grease.

3

Methods

3.1 Field TMR-Dextran Uptake

All steps are performed at room temperature (see Note 6). 1. Take a coverslip with primary neurons attached, and replace the culture medium with the appropriate imaging buffer. Allow the cells to equilibrate for a minimum of 10 min (see Note 7).

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2. Dilute the TMR-dextran in imaging buffer to 50 μM ready for use. You will only need sufficient diluted TMR-dextran solution to fill the imaging chamber-approximately 230 μl. 3. Assemble the imaging chamber (see Note 8), but do not seal the chamber at the top using a coverslip or the clamp from the chamber platform. Set the stimulation settings on the stimulator (40 Hz, 10 s, is sufficient to activate ADBE [5]), and connect to the imaging chamber. 4. Add the TMR-dextran solution to the cells and stimulate immediately (see Note 9). 5. Immediately following termination of stimulation, aspirate the TMR-dextran solution (with either a pipette or vacuum pump), and wash the coverslip twice with imaging buffer (see Note 10). 6. Finish assembling the imaging chamber with the top coverslip and platform clamp, and mount on the microscope. Connect to the perfusion apparatus, and continuously wash the coverslip for 2–5 min with imaging buffer to ensure that all of the non-internalized TMR-dextran has been removed (see Note 11 for an adapted version of the protocol using high-potassium solution to stimulate internalization of TMR-dextran). 7. Focus on the neurons using bright-field illumination, and then take approximately 10 representative fields of as equal neuronal density as possible in both the bright-field (for future reference) and the rhodamine fluorescent channel for TMR-dextran uptake (see Notes 12–14). 8. If Z-stacks were taken to image the TMR-dextran, first combine those images using the Z-projection tool in Fiji to generate one image (see Note 15). 9. The images should then be thresholded to remove any background fluorescence that is not due to TMR-dextran. The remaining TMR-dextran puncta can be automatically counted using the Analyze Particles tool in Fiji (see Note 16). The fields from the same coverslip are then averaged to determine the overall TMR-dextran uptake from that coverslip (see Fig. 2 for example of data generated with this method). 10. An identical process should be undertaken for unstimulated controls. This is employed to calculate the average background signal, which is subtracted from the evoked signal (see Note 17). 3.2 Individual Neuron TMR-Dextran Uptake

1. Perform the protocol above for field TMR-dextran uptake from steps 1 to 6, but using neurons that have been previously transfected with a marker and/or your plasmid of interest (see Notes 18 and 19).

Fig. 2 Example data using the field tetramethylrhodamine (TMR)-dextran uptake assay. Cerebellar granule neurons (CGNs) were stimulated at 40 Hz for 10 s in the presence of 50 μM TMR-dextran. The non-internalized TMR-dextran was immediately washed off and the resulting uptake imaged. (A) Representative images from CGNs either stimulated at 40 Hz for 10 s (left-hand panels) or mock stimulated (right-hand panels). TMR-dextran uptake in nerve terminals is visualized as bright puncta (upper panels). Mock stimulated panels show background fluorescence likely from autofluorescence and cellular debris (which is generally dimmer and does not form discrete puncta). There can be a contribution from non-specific adherence of TMR-dextran. Taken together, background typically represents about 10–20% of the evoked uptake. The lower panels illustrate the thresholded images used for the analysis. Note the difference in the size and shape of the thresholded areas. In the stimulated panel, discrete lines of small puncta representative of TMR-dextran uptake along the neurites are visible, compared to larger aggregate areas present in the unstimulated panel. The range settings for the automated particle counter tool have to be set with respect to the individual microscope to allow accurate and appropriate quantification of the number of TMR-dextran positive nerve terminals. (B) Zoomed areas of panel A shown in the red boxes. (C) Quantification of the presented whole field thresholded stimulated image (Stim) and mock stimulated image (Mock) shown in (A) using the Analyze Particles tool in Fiji with the following settings: size 4–8 pixels and circularity 0–1. Images taken at a magnification of
20. Scale bar ¼ 30 μm

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Fig. 3 Example data using the individual neuron tetramethylrhodamine (TMR)-dextran uptake assay. Cerebellar granule neurons (CGNs) were transfected with an mCerulean (mCer) vector 72 h prior to incubation with 50 μM TMR-dextran solution and stimulation at 40 Hz for 10 s. Non-internalized TMR-dextran was washed away immediately before imaging the neurons for the presence of TMR-dextran containing nerve terminals. (A) Neurite transfected with mCer. (B) The same field, showing TMR-dextran uptake. (C) A merged image of the mCer image and the image showing TMR-dextran uptake. Arrows represent nerve terminals labelled with TMR-dextran, indicating they have undergone activity-dependent bulk endocytosis (ADBE). (D) The axon trace image generated using the Simple Neurite Tracer plug-in in Fiji to quantify the length of neurite. Images taken at a magnification of
40. Scale bar ¼ 5 μm

2. Following mounting of the coverslip, locate a field with transfected neurons, and take an image in both channels to record the position of the transfected neurites as well as the TMR-dextran uptake in the field. Repeat until multiple fields have been imaged (see Note 20). 3. As with the field TMR-dextran uptake, include non-stimulated controls in the experiment design. These should be conducted in exactly the same manner with the same incubation time with the TMR-dextran solution as with the stimulated coverslips. 4. To analyze TMR-dextran uptake, first overlay the transfected neurite image on the TMR-dextran uptake image using Fiji (see Note 21). Manually mark the TMR-dextran puncta which overlay with the transfected neurite in the ROI manager, and record the number of puncta per transfected neurite in the field. 5. Calculate TMR-dextran uptake per length of neurite by measuring the length of the neurite using a tool such as the Simple Neurite Tracer in Fiji (https://imagej.net/Simple_Neurite_ Tracer) and dividing the TMR-dextran uptake for that field by this (see Fig. 3 for example of data generated with this method).

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6. Repeat the analysis for the non-stimulated controls, and subtract the nonspecific uptake to calculate the evoked TMR-dextran uptake.

4

Notes 1. Electrical field stimulation is the most physiological method to stimulate cells; however, it is possible to use a modified version of this protocol that uses high KCl solution (see Note 11). This evokes a clamped depolarization to initiate exocytosis and subsequent ADBE that is equivalent to a field stimulus of 80 Hz for 10 s [11, 14]. 2. An inverted microscope is required so that the oil objective can be in contact with the bottom of the sealed imaging chamber. A high NA objective is required to efficiently detect the dim TMR-dextran puncta. For single neurite experiments, a minimum
40 objective is recommended to ensure sufficient resolution to identify TMR-dextran uptake from individual neurites. 3. For example, if using a Warner imaging chamber, coverslips smaller than 25 mm in diameter will not completely seal the imaging chamber leading to leakage, and larger than 25 mm in diameter coverslips will not fit the platform into which the imaging chamber is subsequently placed. 4. A concentrated (ten times) stock of the imaging buffer can be prepared and stored at 20  C. On the day, an aliquot can be thawed, diluted, and warmed to room temperature for imaging. 5. TMR-dextran is sensitive to both light and temperature. It is delivered as a lyophilized powder for immediate storage at 20  C. Before reconstituting, allow to warm briefly at room temperature. It is recommended to put the appropriate volume of water into the packaging and gently reconstitute using a pipette set to approximately a fifth of the total volume (to ensure the TMR-dextran is completely dissolved and no clumps remain). A failure to reconstitute completely will lead to large fluorescent aggregates being present during imaging. Once in solution, aliquot and store at 20  C until required. Once an aliquot is thawed for use, it should be kept on ice in the dark until dilution into imaging buffer for incubation with the cells. 6. The experiment could be conducted at 37  C, although the loading of the TMR-dextran would have to be performed inside a temperature-controlled chamber.

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7. The equilibration step at the start of the experiment is essential if using cerebellar granule neurons, since these neurons are grown in depolarizing media and require to be repolarized before the experiment. 8. To ensure a good electrical contact at the start of each imaging day, the platinum wires should be gently rubbed with some fine sandpaper to remove any salt deposit. If vacuum grease is required to assemble the imaging chamber, for example, a Warner imaging chamber, the wires should also be cleaned carefully to ensure that they are free from grease. During assembly, vacuum grease should be applied with just sufficient to seal the coverslips. An excess of vacuum grease can squeeze into the chamber, potentially blocking the perfusion tubing exits on the chamber as well as preventing the platinum wires efficiently conducting the stimulation. Excess of vacuum grease in the chamber also provides a surface for nonspecific binding of TMR-dextran which will introduce high background levels of fluorescence into the experiment. 9. It is important to also include non-stimulated coverslips in order to control for any non-specific binding or uptake from the neurons. The coverslip should be set up in exactly the same way as for a stimulated coverslip and the TMR-dextran incubated with the cells for the same period of time as the simulated cells, for example, 10 s to be equivalent to a 40 Hz 10 s stimulus. These coverslips can be used for background subtraction during the analysis step. 10. Directly aspirating the TMR-dextran solution from the coverslip is the most effective way to remove the majority of the non-internalized TMR-dextran quickly, ensuring uptake reflects stimulus-evoked uptake. 11. Adapted protocol using high-potassium solution instead of electrical stimulation: Dilute the TMR-dextran in the high KCl imaging buffer instead of the NaCl imaging buffer, and use this solution in place of the field stimulation. Wash in the same manner as for the field stimulation protocol and assemble the imaging chamber. 12. The output of the field TMR-dextran uptake assay is reliant on the density of the field of neurites; for example, a dense field will have more neurites and thus likely more TMR-dextran uptake. Therefore, it is imperative to image fields of as equal density as possible to ensure comparable results between coverslips and preparations. To control for this, it is possible to preincubate cultures with fluorescent-tagged antibodies that recognize lumenal epitopes of SV proteins, such as synaptotagmin-1 [15]. This will provide an estimation of the

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density of nerve terminals per field, which can be used to normalize the TMR-dextran signal. 13. It can be beneficial to take a small Z-stack of images in the TMR-dextran channel, either manually or automatically, to capture the entire TMR-dextran uptake in a field that may be in different planes of focus. These images can then be combined together at the analysis stage. 14. During the course of the image acquisition, the background can become increasingly bright and diffuse. This is almost certainly due to clumps of TMR-dextran that had adhered to the coverslip, or TMR-dextran solution that had adhered to the edges of the chamber, leaching fluorescence into the field of view. This can be minimized by periodically switching the perfusion on and rewashing the coverslip or by imaging with continuous perfusion. The latter option will use a large volume of buffer, however. 15. The TMR-dextran assay is binary in terms of its readout. Therefore, it reports whether or not a given nerve terminal has internalized TMR-dextran. It is not possible to extract information on the extent of TMR-dextran uptake per nerve terminal (e.g., the amount of fluorescence intensity). Thus, using a maximum fluorescence Z-projection algorithm to compress a Z-stack of images increases the ability of a macro to accurately identify the TMR-dextran puncta without compromising the image data because it minimizes the background fluorescence and maximizes the TMR-dextran signal. 16. A macro to automatically process the images requires thresholding of the individual images, and then to count the particles of the correct size that represent a nerve terminal (approximately 1–3.5 μm2). These values will depend on the microscope and the magnification of the objective and should be optimized by the user. This is important to ensure the accuracy of the counting process. If the range includes particles that are too small to be nerve terminals, or too large representing particles that could not be internalized, the macro will overestimate the count. Conversely, if the range is too stringent, the macro will underestimate the number of nerve terminals accumulating TMR-dextran. 17. The background signal can vary between preparations of cells and batches of dextran but is typically 10–20% of the evoked signal. 18. The density of the cultures is still important for analysis of TMR-dextran uptake in individual neurons because if the cultures are too dense, it is difficult to resolve TMR-dextran internalization in the transfected neurite against a background of uptake in a field of untransfected neurites.

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19. To minimize bleed-through between the channels, when transfecting with a fluorescent marker, maximize the spectral difference between the rhodamine emission of the TMR-dextran and the label of interest; for example, mCerulean is better than GFP. Far-red reporters (such as Alexa-647 or cypHer) are also compatible partners for TMR-dextran imaging [15]. 20. The analysis for this method can be time consuming; therefore, determine in advance the number of fields required for the analysis, bearing in mind the values are combined to provide a coverslip average. 21. There are several ways to do this, but overlaying the images using the Channels Tool in Fiji (Menu: Image ! Color ! Channels Tool) or a plug-in such as the Align RGB plug-in (https://imagej.net/Align_RGB_planes) is effective because it allows toggling of the transfected neurite channel to check that the TMR-dextran puncta are located on the neurite in question. Using a method which permanently overlays the channels can make identification of the TMR-dextran more difficult depending on the fluorescence intensity of the transfection marker.

Acknowledgments This work was supported by grants awarded by Cure Huntington’s Disease Initiative (A-11210 and A-14021; to KS and MC) and the Wellcome Trust (204954/Z/16/Z to MC). References 1. Watanabe S, Rost BR, Camacho-Perez M, Davis MW, Sohl-Kielczynski B, Rosenmund C, Jorgensen EM (2013) Ultrafast endocytosis at mouse hippocampal synapses. Nature 504:242–247 2. Watanabe S, Trimbuch T, Camacho-Perez M, Rost BR, Brokowski B, Sohl-Kielczynski B, Felies A, Davis MW, Rosenmund C, Jorgensen EM (2014) Clathrin regenerates synaptic vesicles from endosomes. Nature 515:228–233 3. Soykan T, Kaempf N, Sakaba T, Vollweiter D, Goerdeler F, Puchkov D, Kononenko NL, Haucke V (2017) Synaptic vesicle endocytosis occurs on multiple timescales and is mediated by Formin-dependent actin assembly. Neuron 93:854–866 4. Granseth B, Odermatt B, Royle SJ, Lagnado L (2006) Clathrin-mediated endocytosis is the dominant mechanism of vesicle retrieval at hippocampal synapses. Neuron 51:773–786

5. Clayton EL, Evans GJ, Cousin MA (2008) Bulk synaptic vesicle endocytosis is rapidly triggered during strong stimulation. J Neurosci 28:6627–6632 6. Kokotos AC, Cousin MA (2015) Synaptic vesicle generation from central nerve terminal endosomes. Traffic 16:229–240 7. Kokotos AC, Peltier J, Davenport EC, Trost M, Cousin MA (2018) Activitydependent bulk endocytosis proteome reveals a key presynaptic role for the monomeric GTPase Rab11. Proc Natl Acad Sci U S A 115:E10177–E10186 8. Cousin MA (2008) Use of FM1-43 and other derivatives to investigate neuronal function. Curr Protoc Neurosci. Chapter 2:Unit 2 6 9. Kavalali ET, Jorgensen EM (2014) Visualizing presynaptic function. Nat Neurosci 17:10–16 10. Clayton EL, Cousin MA (2009) Quantitative monitoring of activity-dependent bulk

Monitoring ADBE Using Fluorescent Dextrans endocytosis of synaptic vesicle membrane by fluorescent dextran imaging. J Neurosci Methods 185:76–81 11. Clayton EL, Anggono V, Smillie KJ, Chau N, Robinson PJ, Cousin MA (2009) The phospho-dependent dynamin-syndapin interaction triggers activity-dependent bulk endocytosis of synaptic vesicles. J Neurosci 29:7706–7717 12. Clayton EL, Sue N, Smillie KJ, O’Leary T, Bache N, Cheung G, Cole AR, Wyllie DJ, Sutherland C, Robinson PJ, Cousin MA (2010) Dynamin I phosphorylation by GSK3 controls activity-dependent bulk endocytosis of synaptic vesicles. Nat Neurosci 13:845–851

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13. Smillie KJ, Pawson J, Perkins EM, Jackson M, Cousin MA (2013) Control of synaptic vesicle endocytosis by an extracellular signalling molecule. Nat Commun 4:2394 14. Cousin MA, Evans GJ (2011) Activation of silent and weak synapses by cAMP-dependent protein kinase in cultured cerebellar granule neurons. J Physiol 589:1943–1955 15. Wenzel EM, Morton A, Ebert K, Welzel O, Kornhuber J, Cousin MA, Groemer TW (2012) Key physiological parameters dictate triggering of activity-dependent bulk endocytosis in hippocampal synapses. PLoS One 7: e38188

Part II Exocytosis

Chapter 8 Quantitative Flow Cytometry-Based Assays for Measuring Constitutive Secretion David E. Gordon, Amber S. Shun-Shion, Asral W. Asnawi, and Andrew A. Peden Abstract Constitutive secretion is predominantly measured by collecting the media from cells and performing platebased assays. This approach is particularly sensitive to changes in cell number, and a significant amount of effort has to be spent to overcome this. We have developed a panel of quantitative flow cytometry-based assays and reporter cell lines that can be used to measure constitutive secretion. These assays are insensitive to changes in cell number making them very robust and well suited to functional genomic and chemical screens. Here, we outline the key steps involved in generating and using these assays for studying constitutive secretion. Key words Constitutive secretion, Flow cytometry, VSV-G, GPI, hGH, Transport, Assay, Trafficking, Biosynthetic transport

1

Introduction Constitutive secretion is a fundamental cellular process required for the delivery of newly synthesized proteins and lipids to the plasma membrane, as well as the exocytosis of extracellular factors such as cytokines and antibodies. To study this process, we have been developing novel quantitative assays for measuring constitutive secretion that are compatible with genetic and chemical screens. The system we have developed is based on a technology originally developed by Ariad Pharmaceuticals to chemically regulate the secretion of recombinant insulin and growth hormone. In this system, the protein to be secreted is tagged with four dimerization domains (originally termed FM4 or CAD domains) that when transfected into cells cause the protein to form large multimers in the endoplasmic reticulum (ER) which are too large to be transported [1, 2]. When the transfected cells are incubated with AP21988 or D/D solubilizer, the multimers rapidly disassemble

Florence Niedergang, Nicolas Vitale and Ste´phane Gasman (eds.), Exocytosis and Endocytosis: Methods and Protocols, Methods in Molecular Biology, vol. 2233, https://doi.org/10.1007/978-1-0716-1044-2_8, © Springer Science+Business Media, LLC, part of Springer Nature 2021

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and are secreted (Fig. 1a). While investigating this system, we observed that the fluorescence or the transfected cells decreased over time as they secrete the reporter. Thus, we thought that this approach could become the basis of a simple flow cytometry-based assay for measuring secretion which would be insensitive to changes in cell number. To generate a robust assay compatible with flow cytometry, we added a GFP tag to the reporter construct, moved it into a γ-retroviral expression system, and generated clonal cell line A)

B)

- Rapamycin/ DD Solubilizer

+ Rapamycin/ DD Solubilizer

1

FCS hGH eGFP DD DD DD DD

1

757aa VSV-G eGFP DD DD DD DD

Golgi

Golgi

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743aa GPI eGFP DD DD DD DD

ER

ER

902aa

Secretory vesicles

Secretory vesicles

Nucleus

Nucleus

Cargo

C)

Cargo

Expected results

Work flow

Viral transduction of reporter construct (4 days) Antibiotic selection (1-2 week) Single cell deposition and clone expansion (2-4 weeks) microscopy (2-4 hours) 2-4 hours)

100 GFP-positive clones 40 clones with uniform expression 10 clones with efficent secretion 2-5 clones with optimal secretion kinetcs

Fig. 1 Overview of the regulated secretion system and steps involved in making reporter cell lines. (a) Diagrammatic representation of the secretory reporter constructs used in this study. All of the constructs share a signal peptide for translocation into the endoplasmic reticulum (ER), eGFP for optical detection, and four dimerization domains (DD) that trap the reporter in the ER. The human Growth Hormone construct (hGH) also has a furin cleavage site after the dimerization domains, so its transport to the TGN can be assessed by proteolytic processing. The vesicular stomatitis virus G protein (VSV-G) reporter construct was generated by cloning the VSV-G transmembrane domain after the dimerization domains. The glycosylphosphatidylinositol (GPI) reporter construct was generated by cloning the GPI signal of human CD55 after the dimerization domains. Numbers indicate amino acids. (b) The secretory reporters when transfected into cells dimerize and form large multimers which are too large to be secreted from the ER. The addition of rapamycin or D/D solubilizer causes the multimers to disassociate and become available for transport from the ER. This process is very rapid, and the reporter constructs can be detected at the cell surface or in the media approximately 20 min after addition of D/D solubilizer. (c) An overview of the main steps involved in generating clonal reporter cell lines and the expected results

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Table 1 Current secretory reporter lines Name of reporter line

Organism and tissue cell lines were derived from

HeLaM_hGH (C1 cells)

Human Cervix

This cell line works very well for RNAi and is able to secrete efficiently in suspension. When the clone is grown adherently, it must be removed from the substrate using a long trypsinization step. For unknown reasons, some of the secreted cargo appears to accumulate at the junctions between cells so making imaging more difficult.

RPE1_hGH (C9 cells)

Human Eye (retinal epithelium)

This cell line shows signs of polarized secretion to the leading edge of the cell. The thinness of the cell makes it very good for live cell imaging experiments. We have found this cell lines to be more difficult to transfect than HeLa-M cells for both RNAi and plasmids. This cell line is unable to secrete in suspension.

Properties of cells

HeLa-M_GPI Human Cervix

This cell line requires a long trypsinization step to remove it from the substrate. The GFP signal in these cells is relatively low so not optimal for live cell imaging. The reporter construct is endocytosed from the cell surface, so long time courses should be avoided. We have not determined if this clone is capable of secreting in suspension.

HeLaM_VSVG

Human Cervix

This cell line requires a long trypsinization step to remove it from the substrate. The GFP signal in these cells is relatively low so not optimal for live cell imaging. The reporter construct is endocytosed from the cell surface, so long time courses should be avoided. We have not determined if this clone is capable of secreting in suspension.

CHO_hGH

Hamster Ovary

These cells are relatively small but have a nice morphology for imaging. We have not determined if this clone is capable of secreting in suspension.

Dmel_hGH (C3 cells)

Drosophila Embryo

These cells are very loosely attached to the substrate and can be easily detached with pipetting, making their harvesting at the end of the experiment very straightforward. Cells are amenable to transient transfection and RNAi.

with stable and uniform expression [3]. This reporter line has proven to be very useful, and many labs are using this system [4– 11]. Since developing this reporter line, we have generated several new cell lines which are derived from different species and cell types increasing the utility of the system (see Table 1) [12]. In addition, we have also developed two new secretory reporters that anchor the

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secreted protein to the membrane either via a transmembrane domain or GPI anchor allowing for cargo-specific trafficking to be studied (Fig. 1b). This chapter will go through the key steps involved in generating, validating, and using these reporter cell lines for measuring constitutive secretion by flow cytometry.

2

Materials

2.1 γ-Retroviral Transfer Plasmids

1. pQCXIP_S-eGFP-DD4-hGH: selection ampicillin and puromycin (see Note 1). 2. pLXIN_S-eGFP-DD4-VSV-G: selection ampicillin and G418. 3. pLXIN_S-eGFP-DD4-GPI: selection ampicillin and G418.

2.2

Cells

1. HeLa-M cells are grown in high-glucose DMEM supplemented with 10% fetal calf serum, 100 IU/mL penicillin, 100 μg/ mL streptomycin, and 2 mM glutamine at 37  C in a 5% CO2 humidified incubator. 2. RPE1 cells are grown in DMEM:F12 (50:50) supplemented with 10% fetal calf serum, 100 IU/mL penicillin, 100 μg/mL streptomycin, and 2 mM glutamine at 37  C in a 5% CO2 humidified incubator. 3. Phoenix retroviral packaging cells are grown in high-glucose DMEM supplemented with 10% fetal calf serum, 100 IU/mL penicillin, 100 μg/mL streptomycin, and 2 mM glutamine at 37  C in a 5% CO2 humidified incubator. The phoenix, γ-retroviral, packaging cells were originally obtained from the Nolan laboratory (Stanford University) but can now be obtained from various commercial suppliers. 4. hGH-based reporter cell lines are maintained using 1 μM puromycin. VSV-G and GPI reporter cell lines are maintained using 0.5 mg/mL G418 (see Note 2).

2.3 Ligands for Inducing Secretion

1. D/D solubilizer (Clontech) stock solution 0.5 mM and working concentration 1 μM (see Note 3). 2. Rapamycin stock solution 1 mM and working concentration 1 μM.

2.4

Equipment

1. Fluorescence-activated cell sorter equipped with a 488 nm laser and 96-well plate autosampler. 2. Flow cytometer (analyzer) equipped with 488 and 647 nm lasers. 3. Inverted fluorescence microscope equipped with a light source that can excite GFP.

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4. A 37  C and 32  C tissue culture incubator. 5. Low-speed centrifuge capable of spinning 15 mL tubes and 96-well plates.

3

Methods The following sections outline the main steps involved in developing and testing reporter cell lines which are suitable for measuring constitutive secretion by flow cytometry.

3.1 Viral Transduction of Secretory Reporter Constructs

The γ-retroviruses generated in this protocol if used inappropriately have the potential to infect the users. Thus, it is important that the appropriate safety measures are put in place before performing this protocol [13] (see Note 4). Please seek advice from the institutional safety committee before performing this work. 1. Day 1, seed approximately five million phoenix cells onto a 10 cm plate (8 mL of media), and allow to adhere for 4 h. 2. Add 21 μL of transfection reagent (TransIT-293) to 200 μL OptiMEM in a 1.5 mL tube, and vortex for 30 s. 3. Incubate for 5 min at room temperature. 4. Add 7 μg of the secretory reporter transfer plasmid, and vortex for 30 s. 5. Incubate for 15 min at room temperature. 6. Add transfection mix dropwise onto the cells and swirl plate to mix. 7. Incubate cells at 37  C overnight. 8. Day 2, remove the media from the phoenix cells, replace with 12 mL of fresh media (appropriate to the cell line to be transduced), and incubate at 32  C overnight (see Notes 5 and 6). 9. Day 3, seed 200,000 cells per mL into a 6-well plate, and allow to adhere for 4 h. 10. Collect the virus-containing media from the phoenix cells using a 20 mL syringe, and mix with polybrene (5 μg/mL final concentration) (see Notes 6 and 7). 11. Filter the virus-containing media using a 0.45 μm filter. 12. Aspirate the media from the plate of cells to be infected, and add the viral supernatant to the cells (2 mL/well). 13. Seal the plate using parafilm, and spin at 2500 rpm (1140  g) for 60 min at room temperature (see Note 8). 14. Remove the parafilm from the plate and incubate at 37  C overnight.

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Fig. 2 Generating and validating the RPEI_hGH secretory reporter line. (a) RPE1 cells were stably transduced with the hGH-based reporter virus and selected using 1 μM puromycin for 1–2 weeks. The selected cells were analyzed using flow cytometry. Representative histograms showing the non-transfected control (gray) and

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15. Day 4, remove the media, trypsinize the cells, resuspend in 30 mL of media, and transfer into a 15 cm diameter plate. Select the cells using the appropriate antibiotics for at least 1–2 weeks (see Notes 9 and 10). After the selection, more than 98% of the cells should be expressing the reporter construct. However, the expression will be heterogeneous (Fig. 2a). To address this issue, clonal cell lines must be generated (see Subheading 3.2). 3.2 Generating Clonal Secretory Reporter Lines

We have observed that the secretion assay works most effectively when performed using clonal cell lines. We have found fluorescentactivated cell sorting (FACS) to be the most efficient and reliable method to generate clonal cells. This process can take between 2 and 4 weeks depending on the rate at which the cells grow. The amount of tissue culture time required to generate clonal cell lines is significant, so it is advisable to only attempt one reporter line at a time (see Fig. 1c for an example of the numbers of clones required to be screened to generate an optimal reporter line). 1. Trypsinize the virally transduced cells, and resuspend in media at one million cells per mL and place on ice (see Note 11). 2. Trypsinize parental cells (non-transduced), and resuspend in 1 mL of media (1 million cells per mL) and place on ice. These are the negative control required for setting up the sorter. 3. Filter the cells using a cell strainer (35 μm nylon mesh) to remove any clumps. 4. Add 200 μL of media to each well of a 96-well flat bottom tissue culture plate (prepare four plates) (see Note 12). 5. Take your sample to your institutional flow cytometry facility, and perform the autocloning (also called single-cell

ä Fig. 2 (continued) transduced cells (blue). (b) GFP-positive cells were autocloned into 96-well plates using flow cytometry and allowed to grow for 2–3 weeks. GFP-positive clones were identified using an inverted fluorescence microscope and expanded into 12-well plates. Positive clones were then analyzed using flow cytometry to establish the uniformity of expression. Representative histograms for several clones showing non-transfected control (gray) and transduced cells (blue). (c) Clones with uniform expression were then assayed for their ability to secrete the reporter construct. Cells were incubated with 1 μM D/D solubilizer for 80 min at 37  C and fluorescence measured using flow cytometry. Representative histograms for several clones showing non-transfected control (gray), cells treated with D/D solubilizer (red), and no treatment (blue). In some instances, the clones are not sensitive to D/D solubilizer and fail to secrete the reporter, so the fluorescence of the clones does not decrease. (d) Transport kinetics for the best RPE1_hGH clone were determined by incubating the cells for the indicated times with 1 μM D/D solubilizer at 37  C and the mean fluorescence determined by flow cytometry. The amount of cargo remaining in the cells after the addition of AP21998 was calculated as a ratio between the control sample (no AP21998) and the experimental samples (+AP21998). The kinetics of transport for the RPE1_hGH clone were plotted in relation to representative HelaM_hGH and Dmel_hGH data

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deposition). Use the non-transduced cells as the negative control (see Note 13). 6. Return the 96-well plates to the 37  C incubator. 7. After 1 week, check for cell growth and contamination. Aspirate any wells that show evidence of bacterial or fungal contamination (see Note 14). 8. After 2 weeks, screen the plate using an inverted fluorescence microscope, and mark the position of the GFP-positive wells using a pen (see Note 15). 9. Once the positive clones have achieved a significant size 2–3 mm, trypsinize and transfer into a 48-well plate (see Note 16). 10. Check the cells every few days, and expand into 24-, 12-, and finally 6-well plates. 11. Once the cells have reached the 12- or 6-well stage, check the stability and clonality of the cells using flow cytometry. Figure 2b shows representative data for four clones isolated when generating the REP1_hGH reporter line. Clones that do not have a uniform expression should be discarded. 12. Determine the ability of the clones to secrete the reporter construct (see Subheadings 3.3 and 3.4). 3.3 Validating and Using hGH Secretory Reporter Cell Line

Once the clonal cell lines have been generated, the ability to secrete the reporter must be assessed. For the hGH-based reporter lines, this is relatively simple as the GFP-tagged secretory reporter is lost from the cells. Thus, the fluorescence of the cells will decrease over time which can be measured by flow cytometry (Fig. 2c). 1. Seed each clone in duplicate onto a 12-well plate (100,000 cells per mL), and grow overnight at 37  C (see Note 17). 2. Aspirate media from one well for each clone. 3. Induce secretion by adding 400 μL of pre-warmed media (37  C) containing 1 μM D/D solubilizer, and incubate for 80 min at 37  C in a tissue culture incubator. 4. Stop secretion by aspirating the media from all of the wells of the plate, and add 1 mL of ice-cold PBS to each well. 5. Aspirate the PBS and add 200 μL of ice-cold trypsin to each well, and incubate for 1–2 h on ice or until cells have detached. 6. Add 200 μL of ice-cold media to each well, mix by pipetting, and transfer the entire sample to a tube suitable for flow cytometry (see Note 18). 7. Measure the mean GFP intensity for each sample using flow cytometry.

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8. Calculate the % of GFP secreted from each clone by dividing the GFP mean intensity of treated (+ D/D solubilizer) and untreated cells. A good clone will secrete approximately 80% of the GFP signal in 80 min. Figures 2c and d show representative data for RPE1_hGH reporter clones. 3.4 Validating VSV-G and GPI Secretory Reporter Lines

For the VSV-G and GPI reporter lines, the GFP is not lost from the cell but is delivered to the cell surface where it accumulates over time. Thus, secretion is followed by measuring the surface levels of GFP using an anti-GFP antibody conjugated to Alexa647 and flow cytometry (Figs. 3 and 4).

Fig. 3 Generating and validating the HeLa-M_VSVG reporter line. (a) Clones with uniform expression of the VSV-G reporter were identified as in Fig. 2, and their ability to secrete assessed using flow cytometry. In this assay, the GFP is anchored to the membrane by the VSV-G transmembrane domain so it is not lost from the cell but is retained at the cell surface. Thus, the surface levels of GFP are determined by staining the cells using an anti-GFP antibody conjugated to Alexa647. Cells were incubated with 1 μM D/D solubilizer for 80 min at 37  C, trypsinized for 1 h on ice, and then labelled with an anti-GFP-Alexa647 antibody for 1 h. The amount of surface labelling was measured using flow cytometry. Representative histograms for several clones showing non-transfected control (gray), cells treated with D/D solubilizer (red), and no treatment (blue). As the secreted GFP reporter accumulates on the cell surface, the mean fluorescence of the cells incubated with the D/D solubilizer will be higher than the non-treated cells. (b) The transport kinetics of a selected clone was determined by incubating the cells for the indicated times with 1 μM D/D solubilizer at 37  C and labelling the cell surface GFP with an anti-GFP-Alexa647-labelled antibody. The mean Alexa647 fluorescence intensity was determined using flow cytometry and plotted against time

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Fig. 4 Generating and testing the HeLa-M_GPI reporter line. (a) Clones with uniform expression of the GPI-anchored reporter were identified as in Fig. 2 and the ability to secrete assessed using flow cytometry. In this assay, the GFP is tethered to the membrane by the GPI anchor so it is not lost from the cells but is retained at the cell surface. Thus, the surface levels of GFP are determined by staining the cells using an antiGFP antibody. Cells were incubated with 1 μM D/D solubilizer for 80 min at 37  C, trypsinized for 1 h on ice, and then labelled with an anti-GFP-Alexa647 antibody for 1 h. The amount of surface labelling was then measured using flow cytometry. Representative histograms for several clones showing non-transfected control (gray), cells treated with D/D solubilizer (red), and no treatment (blue). As the secreted GFP reporter accumulates on the cell surface, the mean fluorescence of the cells will be higher in the cells incubated with the D/D solubilizer. (b) To validate the GPI reporter assay, cells were incubated with a known inhibitor of secretion Brefeldin A and PI-PLC, an enzyme which can cleave the GPI anchor and release the GFP from the cells surface. The cells were incubated with 1 μM D/D solubilizer for 80 min at 37  C in the presence or absence of 2 μM Brefeldin A. The cells were trypsinized for 1 h on ice and labelled with an anti-GFP-Alexa647 antibody for 1 h. In a parallel experiment, the cells were incubated with 1 μM D/D solubilizer for 80 min at 37  C. The cells were incubated with PI-PLC (1 unit) for 1 h on ice and trypsinized and labelled as above. The amount of surface labelling was measured using flow cytometry and normalized to the untreated sample. (c) To determine if the GPI reporter requires SNARE proteins for its transport, we depleted the ER-localized SNARE syntaxin 5 (STX5) and its Sec-Munc protein Sly1 using RNAi [3]. The cells were incubated with 1 μM D/D solubilizer for 80 min at 37  C, trypsinized for 1 h on ice, and labelled with an anti-GFP-Alexa647 antibody for 1 h. The amount of surface labelling was then measured using flow cytometry and normalized to the mocktreated sample

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1. Seed each clone in duplicate onto a 12-well plate (100,000 cells per mL), and grow overnight (see Note 19). 2. Aspirate media from one well per clone. 3. Induce secretion by adding 400 μL of pre-warmed media (37  C) containing 1 μM D/D solubilizer, and incubate for 80 min at 37  C in a tissue culture incubator (see Note 20). 4. To stop secretion, aspirate the media from all of the wells, and add 1 mL of ice-cold PBS. 5. Aspirate the PBS and add 200 μL of ice-cold trypsin to each well, and incubate for 1–2 h on ice or until the cells have detached. 6. Quench the trypsin using 400 μL of ice-cold media, and spin down the cells at 1000 rpm for 5 min (see Notes 21 and 22). 7. Remove the media, resuspend cells in 200 μL of ice-cold media containing the anti-GFP-Alexa647-labelled antibody, and incubate for 1 h on ice (1 μL of antibody per 200 μL). 8. Spin down the cells at 1000 rpm for 5 min. 9. Wash the cells by resuspending in 400 μL of ice-cold media. 10. Repeat steps 6 and 7 two more times. 11. Measure the mean Alexa647 intensity for each sample using flow cytometry. If the cells have been treated with RNAi or chemical inhibitors, the intensity of the cell surface signal can be normalized to the untreated or mock samples (see Fig. 4b and c for representative data).

4

Notes 1. The secretory reporter constructs generated by our group are based on the pC4S1-FM4-FCS-hGH plasmid included in the RPD Regulated Secretion/Aggregation Kit (ARIAD Pharmaceuticals). This plasmid is no longer available from ARIAD Pharmaceuticals but can be obtained from Clontech (vector renamed prHom-Sec1 and the FM4 domains renamed DmrD). We modified this construct by inserting eGFP (Clontech) after the signal peptide using a unique XbaI site. We have renamed the FM4 domains dimerization domains (DD) in all of our constructs. The reporter was then cloned into the retroviral transfer plasmid pQCXIP (Clontech). The VSV-G based reporter was generated by excising hGH from pC4S1-FM4FCS-hGH (SpeI/BamHI) and replacing it with the transmembrane domain of VSV-G. We initially cloned this reporter into pQCXIP. However, we discovered that the construct was expressed at very high levels causing a significant amount of the reporter to leak onto the cell surface in the absence of the

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ligand used to induce secretion. Thus, we cloned the reporter into pLXIN which is a retroviral transfer plasmid with a significantly weaker promoter. The GPI construct was generated in a similar manner as the VSV-G reporter, and the hGH was replaced with the GPI signal of human CD55 and cloned into pLXIN. All of the transfer plasmids were amplified in DHFα bacteria and purified using a maxiprep kit. We have found that the concentration of the transfer plasmid has a significant impact on viral titer, so one should aim to have a concentration of at least 1 mg/mL. 2. We have found that the reporter cell lines behave consistently over a number of passages (10–20) as long as the cells are maintained in selection. 3. The reporter constructs form large multimers in the ER that are too large to be exported by the intracellular transport machinery. In the original version of our assay, we used the small molecule AP21998, supplied by ARIAD Pharmaceuticals, to induce their dissociation and subsequent secretion. This small molecule is no longer available. However, a similar molecule can be purchased from Clontech (D/D solubilizer) and is effective at 1 μM. We have found that rapamycin is also very effective and can be used at similar concentrations as the D/D solubilizer. However, prolonged incubations should be avoided as this can induce autophagy. 4. The tissue culture media and plasticware used during the viral transduction contain viral particles so should be handled with care. The media and plasticware should be inactivated using bleach or a detergent-based disinfectant in line with your institutional procedures. To avoid aerosols, do not aspirate media from the cells; instead, transfer using a pipette. 5. Take care when adding media to the cells as it is easy to wash them from the plate. 6. We have found that, in most instances, it is not necessary to produce the virus at 32  C. However, we recommend that this step is initially followed to ensure optimal virus titer. 7. Directly pipette the polybrene into the syringe, and pull back the plunger by 1–2 cm. This will make the syringe easier to use. The virus-containing media can be frozen and stored in the 80  C at this stage. However, the titer of the virus will reduce by approximately 50%. 8. Great care should be taken when sealing the plate with parafilm. It is very easy to knock the plate and spill the viruscontaining media. To help reduce the chance of this happening, directly apply pressure to the top of the plate when wrapping in parafilm.

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9. It is important that the cells do not become confluent during the selection process as this protects the cells from the selection reagent. If the cells look like they are becoming confluent, reduce the density of the cells by splitting (1/3 or 1/5). 10. As the secretory reporter constructs contain eGFP, the success of the viral transduction and selection can be monitored using an inverted fluorescence microscope. 11. Only 1–2 million cells are needed for the autocloning so any spare cells should be frozen down at this point. For most immortal cell lines, viability will be greater than 98%, so a viability stain is not normally necessary for the sorting. 12. We have found that it can be useful to maintain the selection drug during the autocloning process. However, we normally reduce the concentration by 25–50%. In some instances, we have found it useful to use conditioned media or increase the amount of fetal calf serum in the media when performing the cloning step. 13. When performing the sorting, we would recommend that the top 50% of GFP-positive cells are gated and used for the autocloning. 14. In some instances, more than 1 cell will be deposited per well. Discard any wells that have more than 1 colony growing in them. 15. We have observed that the growth rate of different clones can vary significantly. We have found that the slow-growing clones do not expand well so are best avoided. 16. To aid identifying the wells during trypsinization, also mark the bottom of the plate using a marker pen. We would normally pick between 5 and 20 clones per plate if the autocloning has been successful. 17. Remember to prepare non-transduced cells (parental cells) as an unstained control for setting up the flow cytometer. 18. When analyzing small volumes by flow cytometry, it is often useful to use 1.2 mL cluster tubes. The sample is directly placed in the cluster tube, and the cluster tube is then inserted inside a larger tube which attaches to the flow cytometer. 19. Once you are happy that the reporter lines are secreting reproducibly, they can then be used for RNAi, CRISPR/Cas9, or chemical inhibitor screens (see Fig. 4b, c for pilot data). 20. To confirm that the increase in Alexa647 signal is due to the delivery of the reporter construct to the cell surface, inhibitors of secretion can be added during this step (2 μM Brefeldin A is an effective inhibitor of this assay; Fig. 4b). In addition, to confirm the specificity of the anti-GFP-Alex647 antibody,

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parental HeLa-M cells can be incubated with the antibody. Any observed signal is due to nonspecific binding of the antibody. It is also worth titrating the antibody to find the appropriate working concentration as this can have a large effect on the reliability of the assay. 21. The specificity of the Alexa647 staining in the HeLa-M_GPI reporter cells can also be tested by incubating the cells with PI-PLC. PI-PLC is an enzyme that specifically cleaves the GPI linkage releasing the secretory reporter from the cell surface (incubate the cells with 1 unit for 30 min on ice). 22. When handling a large number of samples, it can be useful to perform the staining and wash steps in a 96-deep well plate (capacity 1 mL). Once the cells are spun down, the media can be removed by simply flicking or rapidly inverting the plate, thus significantly reducing the time needed for aspirating the samples.

Acknowledgments We would like to thank the MRC (G120/952) and BBSRC (BB/L002841/1 and BB/S009566/1) for supporting this work. A. S. Shun-Shion was funded by the BBSRC White Rose DTP studentship (BB/J014443/1). References 1. Rollins CT, Rivera VM, Woolfson DN, Keenan T, Hatada M, Adams SE, Andrade LJ, Yaeger D, van Schravendijk MR, Holt DA, Gilman M, Clackson T (2000) A ligandreversible dimerization system for controlling protein-protein interactions. Proc Natl Acad Sci U S A 97:7096–7101. https://doi.org/ 10.1073/pnas.100101997 2. Rivera VM, Wang X, Wardwell S, Courage NL, Volchuk A, Keenan T, Holt DA, Gilman M, Orci L, Cerasoli F Jr, Rothman JE, Clackson T (2000) Regulation of protein secretion through controlled aggregation in the endoplasmic reticulum. Science 287:826–830. https://doi.org/10.1126/science.287.5454. 826 3. Gordon DE, Bond LM, Sahlender DA, Peden AA (2010) A targeted siRNA screen to identify SNAREs required for constitutive secretion in mammalian cells. Traffic 11:1191–1204. https://doi.org/10.1111/j.1600-0854.2010. 01087.x 4. Bond LM, Peden AA, Kendrick-Jones J, Sellers JR, Buss F (2011) Myosin VI and its binding

partner optineurin are involved in secretory vesicle fusion at the plasma membrane. Mol Biol Cell 22:54–65. https://doi.org/10. 1091/mbc.E10-06-0553 5. Brandstaetter H, Kruppa AJ, Buss F (2014) Huntingtin is required for ER-to-Golgi transport and for secretory vesicle fusion at the plasma membrane. Dis Model Mech 7:1335–1340. https://doi.org/10.1242/ dmm.017368 6. Lamb CA, Nuhlen S, Judith D, Frith D, Snijders AP, Behrends C, Tooze SA (2016) TBC1D14 regulates autophagy via the TRAPP complex and ATG9 traffic. EMBO J 35:281–301. https://doi.org/10.15252/ embj.201592695 7. Miller CN, Smith EP, Cundiff JA, Knodler LA, Bailey Blackburn J, Lupashin V, Celli J (2017) A Brucella type IV effector targets the COG tethering complex to remodel host secretory traffic and promote intracellular replication. Cell Host Microbe 22(317–329):e7. https:// doi.org/10.1016/j.chom.2017.07.017

Quantitative Flow Cytometry-Based Assays for Measuring Constitutive Secretion 8. Song K, Gras C, Capin G, Gimber N, Lehmann M, Mohd S, Puchkov D, Rodiger M, Wilhelmi I, Daumke O, Schmoranzer J, Schurmann A, Krauss M (2019) A SEPT1-based scaffold is required for Golgi integrity and function. J Cell Sci 132:jcs225557. https://doi.org/10.1242/ jcs.225557 9. Wendler F, Gillingham AK, Sinka R, RosaFerreira C, Gordon DE, Franch-Marro X, Peden AA, Vincent JP, Munro S (2010) A genome-wide RNA interference screen identifies two novel components of the metazoan secretory pathway. EMBO J 29:304–314. https://doi.org/10.1038/emboj.2009.350 10. Winslow AR, Chen CW, Corrochano S, Acevedo-Arozena A, Gordon DE, Peden AA, Lichtenberg M, Menzies FM, Ravikumar B, Imarisio S, Brown S, O’Kane CJ, Rubinsztein DC (2010) Alpha-Synuclein impairs

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macroautophagy: implications for Parkinson’s disease. J Cell Biol 190:1023–1037. https:// doi.org/10.1083/jcb.201003122 11. Wong M, Munro S (2014) Membrane trafficking. The specificity of vesicle traffic to the Golgi is encoded in the golgin coiled-coil proteins. Science 346:1256898. https://doi.org/10. 1126/science.1256898 12. Gordon DE, Chia J, Jayawardena K, Antrobus R, Bard F, Peden AA (2017) VAMP3/Syb and YKT6 are required for the fusion of constitutive secretory carriers with the plasma membrane. PLoS Genet 13: e1006698. https://doi.org/10.1371/journal. pgen.1006698 13. Mosier D (2004) Introduction for “safety considerations for retroviral vectors: a short review”. Appl Biosafety 9:68–75. https://doi. org/10.1177/153567600400900203

Chapter 9 High-Throughput Screening for Insulin Secretion Modulators Michael A. Kalwat Abstract The application of forward chemical genetics to insulin secretion in high-throughput has been uncommon because of high costs and technical challenges. However, with the advancement of secreted luciferase tools, it has become feasible for small laboratories to screen large numbers of compounds for effects on insulin secretion. The purpose of this chapter is to outline the methods involved in high-throughput screening for small molecules that chronically impact pancreatic beta cell function. Attention is given to specific points in the protocol that help to improve the dynamic range and reduce variability in the assay. Using this approach in 384-well format, at least 48 and as many as 144 plates can theoretically be processed per week. This protocol serves as a guideline and can be modified as required for alternate stimulation paradigms and improved upon as new technologies become available. Key words High-throughput screening, Insulin secretion, Gaussia luciferase, Small molecules, Pancreatic beta cells

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Introduction Diabetes afflicts over 425 million people worldwide and results from pancreatic islet dysfunction [1]. Blood glucose homeostasis is largely maintained by the concerted action of insulin secretion from islet beta cells. Consumption of nutrients leads to elevated blood glucose levels which are sensed by beta cells, stimulating them to release insulin. Insulin signals to peripheral skeletal muscle and adipose tissue to take up glucose, restoring euglycemia. In diabetes, the beta cells fail to release sufficient amounts of insulin and eventually fail or die. Chronically elevated blood glucose levels lead to disease pathologies including nephropathy, neuropathy, and retinopathy. The inability of current therapies to preserve or fully restore diabetic islet function is directly tied to insufficient knowledge of nutrient-regulated secretion and limited pharmacological targets and interventions. Compounds that enhance or inhibit

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insulin secretion represent useful agents to identify important β-cell regulatory pathways and provide novel pharmacological opportunities to stabilize β-cell function in disease. However, highthroughput screens for chemical perturbagens of insulin secretion are rare [2–5], largely due to high cost and technical challenges. The goal of this methods chapter is to outline one successful high-throughput screening strategy and point out critical steps and places where the method may be improved upon. This approach relies on using a stable beta-cell line expressing a luciferase reporter that is co-secreted with insulin [4, 5]. This system has been used at the bench in low- to medium-throughput experiments [6, 7]. Others have also used a similar system to discover compounds with short-term effects on glucose-stimulated insulin secretion using beta cells in suspension [4, 8]. However, to discover compounds with more chronic, long-term effects, alternate technical methods are required.

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Materials

2.1 Required Equipment and High-Throughput Screening Facility Capabilities

1. BioMek FX robotic liquid handler or suitable alternative. 2. BioTek Multiflo FX liquid handler and 5 μL cassette (see Note 1). 3. Perkin Elmer EnVision multimode plate reader or equivalent. 4. Tabletop swing bucket centrifuge with collecting trays (see Note 2). 5. Cell culture materials: T175 flasks, opaque white 384-well tissue culture-treated plates.

2.2 InsGLuc MIN6 Cell Culture

1. InsGLuc-MIN6 cells (see Note 3).

2.3 InsGLuc Secretion Assay

1. Stock solutions (see Note 5).

2. InsGLuc-MIN6 cell media: DMEM containing phenol red, 25 mM glucose, 0.11 g/L sodium pyruvate, 15% FBS, 100 U/mL penicillin, 100 μg/mL streptomycin, 292 μg/mL L-glutamine, 50 μM β-mercaptoethanol, 250 μg/mL G418 (see Note 4).

2. Coelenterazine (CTZ): 1 mg/mL (2.36 mM) coelenterazine in acidified methanol. Keep stocks in screw-capped tubes sealed with parafilm at 80  C (see Note 6). 3. Diazoxide: 11.53 mg/mL (50 mM) in 0.1 N NaOH. Freeze stocks at 20  C. 4. 3 M KCl: 26.8 g in 120 mL water. Filter sterilize and store stock at room temperature.

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5. 2 M glucose: 36 g in 100 mL water. Filter sterilize and store stock at 4  C. 6. Krebs-Ringer Bicarbonate Hepes (KRBH) buffer: 5 mM KCl, 120 mM NaCl, 15 mM Hepes pH 7.4, 24 mM NaHCO3, 1 mM MgCl2, and 2 mM CaCl2 (see Note 7). 7. Diazoxide-KRBH: KRBH with 250 μM diazoxide from the stock in 0.1 N NaOH. Add an equal amount of 0.1 N HCl to maintain the pH of the solution. 8. 2 KCl/glucose in diazoxide-KRBH: 70 mM KCl and 40 mM glucose. 9. Ascorbate KRBH: 5 mM KCl, 15 mM Hepes pH 7.4, 24 mM NaHCO3, 1 mM MgCl2, 2 mM CaCl2, and 300 mM sodium ascorbate (see Note 8). 10. Gaussia luciferase working solution: Add 1.5 μL per mL of ascorbate-KRBH to result in 3.54 μM CTZ. Prepared 15 min before use.

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Methods

3.1 Expansion and Plating of InsGLuc MIN6 Cells

1. On day 1 (e.g., Monday), trypsinize 8–10 confluent T175 flasks. Suspend detached cells in media, and spin down in swing bucket centrifuge at 800 rpm for 5 min to remove trypsin. Resuspend cell pellets in 10 mL per flask used, and pass through a 45 μm cell strainer into a fresh 50 mL conical tube. 2. Count the cells, and dilute to 1.5  106 cells/mL in 500 mL of InsGLuc MIN6 cell media (see Note 9). Confirm diluted cell concentration on cell counter. 3. Plate 15–17  106 cells in each of 8–10 new T175 flasks with 20 mL media for plating/screening the following week (see Note 10). 4. Using BioTek Multiflo with the 5 μL cassette, prime at least 15–20 mL through the tubing. Set the liquid handler to dispense 50 μL per well to each of 24,384-well opaque white cell culture dishes (see Note 11). Give the flask a gentle swirl in between each plating to ensure a homogenous cell slurry. Place the dishes without stacking in a 37  C tissue culture incubator for 24 h. 5. On day 2 (after 24 h), the cells should be treated with control and test compounds on a liquid handling robot (see Note 12). The media does not need to be changed. Place the drug-treated cells back into the incubator for 24 h.

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3.2 InsGLuc Secretion Assay

1. On day 3, prepare the buffers needed for the screen: KRBH, Dz-KRBH, 2 KCl/glucose-Dz-KRBH, and Asc-KRBH (see Note 13). 2. Prime the BioTek cassette on the Multiflo with KRBH (see Note 14). 3. Centrifuge plates upside down in collection trays at 30  g for 1 min to remove the media. Wash the plate twice with KRBH buffer, first with 75 μL per well and then with 50 μL per well, centrifuging and blotting on paper towels between each wash. After the final spin, add 25 μL of KRBH-Dz buffer to all wells (see Note 15). 4. Incubate 60 min in tissue culture incubator. Note the time that particular set of plates was placed in the incubator. 5. Prime the BioTek cassette with 2 KCl/glucose Dz-KRBH. After the 60 min incubation, remove the set of plates, and add 25 μL of 2 KCl/glucose Dz-KRBH to all wells. Note the time and return the set of plates to the incubator for 60 min. 6. During the incubation, prepare the Gaussia luciferase working buffer. 7. After the 60 min incubation, remove plates from incubator (see Note 16). Prime the cassette with Gaussia luciferase working solution, and add 20 μL to each well. 8. Stack plates without lids and load into the Perkin Elmer EnVision plate reader (or suitable alternative). Read the plates at 0.1 s integration per well (see Note 17). An example of a 24-plate run is shown in Fig. 1a. The summation of an entire 100,000 compound screen is shown as a plot in Fig. 1b.

4

Notes 1. Carefully calibrate the cassette before starting a screen, and always rinse it well with 70% ethanol. It is helpful to take apart the cassette every few weeks after heavy use and clean the nozzles out. Preferably, two cassettes should be used to reduce wear and tear during screening, one for plating cells and one for performing the washes and stimulations in the assay. 2. Access to two adjacent swing bucket centrifuges facilitates staggering plates in sets of four to allow a smooth 24-plate run. Beckman Coulter reservoirs (Cat # 372784) are a useful option for spinning 384-well plates upside down to collect media or buffer. Centrifuging the media/buffer from plates was chosen subsequent to testing a liquid handler with a 96-tip vacuum manifold. Even on the gentlest setting, the vacuum manifold disrupted the MIN6 cell layer and led to variable

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Fig. 1 Example of high-throughput screening results. (a) A representative example of a 24-plate run (7680 test compounds) with positive controls (low signal) in columns 1 and 24 and negative control DMSO (high signal) in columns 2 and 23. Columns 3 through 22 are all test compounds. Blue is suppressed and red is increased with respect to the negative control columns. (b) Waterfall plot displaying overall results of an entire highthroughput primary screen using the InsGLuc reporter in MIN6 cells. Hits with |Z-score|  3 are intended to be confirmed in triplicate followed by dose-response experiments and follow-up studies. Plot was generated in TIBCO Spotfire

Z-scores. Centrifuging the inverted plates at low speed was gentler, took a similar amount of time, and led to higher and more consistent Z-scores. 3. InsGLuc MIN6 cells will be shared upon request. Others can also regenerate their own stable lines by transfecting the pcDNA3.1+rIns-hIns-eGLuc2.1 plasmid (Addgene #89928) into MIN6 or INS1 cells and selecting with G418. Alternatively, lentivirus can be made from the pLenti-rIns-hInseGLuc2.1 vector (Addgene #89927), and transduced cells can be selected with puromycin. 4. The stable InsGLuc MIN6 cell line was originally selected using G418 and so it is included in the media to help maintain expression of the transgene [5]. The line can be generated using lentivirus and puromycin selection as well [4].

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5. When beginning a large screen, we found it helpful to make large stocks of all buffer components to last through the entire screen. This improves the stability and repeatability of the assay. 6. Acidified methanol is 1.06% HCl in methanol. Coelenterazine activity is highly batch dependent. Anecdotally, during these studies, CTZ from RPI and NanoLight was tested. While the RPI CTZ had lower activity than the NanoLight CTZ, it was used for the screen because the slightly lower signal improved the dynamic range of the assay, leading to better Z-scores. It is recommended to test a small amount and then purchase enough of the same lot of CTZ to last for the entire screen in order to avoid switching lots/batches/vendors in the middle of the screen. 7. Normally, KRBH contains bovine serum albumin (BSA) at 1 mg/mL. We omitted BSA from the buffer to prevent bubbles and drips from occurring on the nozzles of the BioTek Multiflo cassette. 8. Sodium ascorbate is included to increase the stability of the Gaussia luciferase reaction with its substrate coelenterazine [9]. Sodium ascorbate contributes sodium ions, so NaCl is omitted from the buffer. The Ascorbate-KRBH buffer containing coelenterazine will eventually be diluted into KRBH (1:3), reducing the final salt concentration. Coelenterazine can also be added to plain KRBH or PBS; however, the half-life may be shorter. Avoid using any buffer that contains detergents which may lyse the cells and cause release of stored Gaussia luciferase. 9. 1.5  106 cells/mL results in a final concentration of 7.5  104 cells per 50 μL per well of a 384-well plate. This concentration was determined to result in the best Z-scores in this assay. Therefore, 28.8  106 cells per 384-well plate are needed. A confluent T175 yields 120  106 cells. Because at least 7 T150s are needed to plate 24  384 well as well as plate new T175s for the following week, it is suggested to use 8–10 T175s and make >500 mL of cell suspension. This helps ensure there is sufficient volume to prime the BioTek cassette and account for the cassette dead volume. 10. Change media on the flasks on Wednesday and Friday if cells are plated on Monday. For cells plated on Tuesday, media can be changed on Friday only. 11. It is helpful when beginning this assay to also seed a clear bottom 384-well dish to monitor the density of the cells after plating and on the day of the assay, as well as confirm effects of any control compounds that cause visible changes to cell morphology.

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12. MIN6 cells tolerate a 24 h treatment with 1–3% DMSO without impact on secretory response in this assay [5]. Therefore, 0.5 μL of DMSO (100%) or test compounds (0.5 mM) are typically added to the cells in 50 μL of media. The first or last column of the plate can be used for a positive control, such as thapsigargin (100 nM) or any chronic treatment that represses beta-cell function. 13. Diazoxide targets the KATP channel, holding it open. In this state, glucose can only elicit exocytosis in the presence of depolarizing concentrations of KCl [10]. The diazoxide paradigm was chosen because it stimulates a large amount of secretion in response to nutrients without needing other drugs/ hormones like GLP-1 or forskolin/IBMX. This generates a large dynamic range in the secretion assay; however, a caveat is that inhibitors are much easier to detect than activators. To screen more specifically for chronic enhancers of beta-cell function, a less potent stimulus (such as glucose alone) can be used in the assay. In that case, a stimulus like forskolin/IBMX or Dz/KCl/glucose can be used in the positive control column for activation and glucose stimulation alone for compoundtreated wells. The dynamic range may be smaller, but enhanced secretion should be more easily detected. 14. Excess primed buffer can be conserved by removing the nozzle/head portion of the cassette and aiming it into a 25 mL reservoir during the priming. The reserved buffer can be carefully added back to the source bottle. 15. With two centrifuges that can hold four plates each, a set of eight plates can be washed in 15 min. Three sets of eight can be done in under an hour, allowing sufficient time to prime the 2 KCl/glucose Dz-KRBH for stimulating the cells after 1 h of preincubation. This technical aspect is the main reason for choosing 24 plates in a run. Alternate methods or equipment may allow for increased numbers of plates to be screened at once. 16. Because the Gaussia luciferase working solution is at room temperature, cooling the plates to room temperature by placing them on a cool metal surface for 5 min prior to adding the substrate working solution will reduce variability in the luciferase reaction due to temperature changes. 17. Plates can be read within 5–10 min after substrate addition. Luciferase signal is still sufficient 20–30 min later as long as a sensitive reader is used.

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Acknowledgments Thanks to Derk Binns for technical assistance and proofreading this manuscript. Thank you to the UTSW High-Throughput Screening Core facility, especially Shuguang Wei, for helpful discussions and advice. Thank you to Magdalena Grzemska for technical assistance. This work was supported by a Juvenile Diabetes Research Foundation Strategic Research Award 2-SRA-2019-702-Q-R (to MAK). Early parts of the development of this methodology were supported by an F32 DK100113 to Michael A. Kalwat and an R37 DK034128 and Welch (I-1243) to Melanie H. Cobb. The UTSW HTS facility is supported by an NIH NCI grant P30 CA142543. References 1. International Diabetes Federation (2017) IDF Diabetes Atlas, Eighth edition. Brussels, Belgium 2. Wu W et al (2008) Identification of glucosedependent insulin secretion targets in pancreatic beta cells by combining definedmechanism compound library screening and siRNA gene silencing. J Biomol Screen 13 (2):128–134 3. Lee JA et al (2011) Open innovation for phenotypic drug discovery: the PD2 assay panel. J Biomol Screen 16(6):588–602 4. Burns SM et al (2015) High-throughput luminescent reporter of insulin secretion for discovering regulators of pancreatic beta-cell function. Cell Metab 21(1):126–137 5. Kalwat MA et al (2016) Insulin promoterdriven Gaussia luciferase-based insulin secretion biosensor assay for discovery of beta-cell glucose-sensing pathways. ACS Sens 1 (10):1208–1212

6. Kalwat M, Cobb MH (2019) Measuring relative insulin secretion using a co-secreted luciferase surrogate. J Vis Exp 148 7. Kalwat MA et al (2018) Chromomycin A2 potently inhibits glucose-stimulated insulin secretion from pancreatic beta cells. J Gen Physiol 150(12):1747–1757 8. Burns SM, Wagner BK, and Vetere A (2018) Compounds and methods for regulating insulin secretion. International Patent Number: WO2018175324A1 9. Ohmiya Y, Wu C (2010) Stabilizing composition and stabilizing method of coelenterazine solution for high-throughput measurement of luciferase activity. U.S. Patent 7,718,389 B2 10. Henquin JC (2000) Triggering and amplifying pathways of regulation of insulin secretion by glucose. Diabetes 49(11):1751–1760

Chapter 10 Different Approaches to Record Human Sperm Exocytosis Laila Suhaiman, Karina Noel Altamirano, Alfonsina Morales, and Silvia Alejandra Belmonte Abstract Acrosome reaction is an exocytic process that enables a sperm to penetrate the zona pellucida and fertilize an egg. The process involves the fenestration and vesiculation of the sperm plasma membrane and outer acrosomal membrane, releasing the acrosomal content. Given the importance of the acrosome secretion in fertilization, many different methods have been developed to detect the acrosome reaction of sperm. In this chapter, we describe detailed practical procedures to assess the acrosomal status of human spermatozoa. To do this, we resorted to light optical and epifluorescence microscopy, flow cytometry, and transmission electron microscopy. We also itemize the protocol for real-time measurements of the acrosome reaction by confocal microscopy. Further, we discuss the level of complexity, costs, and the reasons why a researcher should choose each technique. This chapter is designed to provide the user with sufficient background to measure acrosomal exocytosis in human sperm. Key words Acrosome exocytosis, Human sperm, Acrosome reaction, Recording sperm secretion

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Introduction Secretion of neurotransmitters, hormones, and enzymes is a fundamental biological activity of the cell and is achieved by vesicular exocytosis, that is, fusion of secretory vesicles with the plasma membrane. Regulated secretion is a central issue for the specific function of many cells; for instance, mammalian sperm exocytosis is essential for egg fertilization. Fertilization is an essential step in sexual reproduction and consists of a carefully orchestrated series of events that culminate with the generation of a genetically unique zygote [1]. Sperm-egg coat penetration, fusion with the egg’s plasma membrane, and finally, fertilization require the release and exposure of cell components resulting from exocytosis of the unique sperm vesicle [2].

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1.1 General Features of a Human Spermatozoon

The spermatozoon is a highly polarized cell constituted by a head and a tail or flagellum. The head externally shows two major domains, the acrosomal region and the post-acrosomal region, separated by the equatorial region. It contains a nucleus and an acrosome, which is a Golgi-derived, lysosome-like, very large electron-dense granule covering about 50% of the nucleus in human sperm [3]. The flagellum contains a 9 + 2 array of microtubules, sheath proteins, and mitochondria, which are spirally arranged in the flagellar midpiece, contributing to power its movement. The head and the tail are joined at the neck. The plasma membrane wraps these structures leaving a little cytoplasm inside. Sperm are incapable of synthesizing proteins or nucleic acids. The only purpose of this terminally differentiated cell is to find, fuse, and deliver their genetic information to the egg (Fig. 1). The large and flat granule covering half of the nucleus, at its apical part, is not just a bag of soluble enzymes, easily released when the outer acrosomal membrane fuses with the plasma membrane. Its content is electron-opaque, given that it contains an important acrosomal matrix required in fertilization. This matrix is a molecular scaffold assembly that is dismantled by a self-regulated mechanism driven, in part, by proteolysis [4]. The acrosome has all the characteristics attributed to secretory vesicles. It undergoes regulated exocytosis when challenged by physiological stimuli, like progesterone or zona pellucida glycoproteins. Although a continuous membrane surrounds the acrosome granule, the membrane consists of different parts that involve distinct functions. The portion overlying the nucleus is termed the inner acrosomal membrane (IAM) and that underlying the plasma membrane is termed the outer acrosomal membrane (OAM). As stated above, the acrosome contains soluble proteins and an insoluble acrosomal matrix, which have their own patterns of release [5, 6]. Soluble factors are readily released once the exocytosis starts, whereas matrix components remain associated for prolonged periods after exocytosis.

1.2 Exocytosis of the Acrosome: Morphological Changes of the Granule

Upon ejaculation in the female genital tract, millions of sperm ascend the uterus. However, only a few pass through the oviduct to reach the ampulla where the fertilization occurs. During this transit, sperm acquire the ability to fertilize eggs through a process defined as capacitation, which consists of a series of physiological and molecular changes that sperm acquire in the female reproductive tract [7]. These changes pertain to the sperm motility pattern, called “hyperactivation,” [8] and to their ability to undergo sperm granule exocytosis. In the proximity of the egg, progesterone and zona pellucida glycoproteins stimulate mammalian sperm to release the contents of the acrosome granule, a key event in fertilization. Acrosomal exocytosis is an all-or-nothing event that involves the opening of multiple fusion pores between the outer acrosomal

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Fig. 1 Top. Scheme showing general features of the human spermatozoa. The major domains on the sperm head are the acrosomal (a), post-acrosomal region (PA), and the equatorial segment (ES). The flagellum can be divided into three regions: the middle piece, containing the helically wrapped mitochondria, the principal piece, and the end piece. The schematic representation of a sagittal section reveals the detailed structure of the head. The plasma membrane (PM) overlies the outer acrosomal membrane (OAM), in turn the inner acrosomal membrane (IAM), overlies the nuclear envelope. Bottom. Diagram of the sequence of events that occurs in spermatozoa’s head during human sperm acrosome reaction. (a) Represents the head of an intact acrosome sperm as described at the top. (b) After a stimulus, the acrosome swells showing deep invaginations of the OAM. The protruding edges of the invaginations tightly appose to the PM. (c) Represents the stage where fusion occurs between the OAM and PM at multiple sites. As exocytosis proceeds, the acrosomal matrix disperses and hybrid vesicles are sloughed. (d) Finally, when the acrosome is lost, the IAM becomes the limiting membrane of the cell and now the spermatozoa has completed its reaction. The acrosomal plasma membrane fuses with the OAM at the equatorial segment and maintains cytoplasmic integrity in the posterior head. The equatorial segment overlies that region where the inner and outer acrosomal membranes merge. The equatorial region of the sperm is ready to fuse with the oocyte at this point

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membrane and the plasma membrane. This Ca2+-regulated exocytosis is known as “sperm acrosome reaction.” Initially, the importance of exocytosis in mammalian fertilization was unclear, but the phenomenon soon became recognized as a prerequisite for fertilization. Acrosome reaction involves the prominent rearrangement of membranes in the sperm head. In resting sperm, the majority of cells show an acrosome with electron-dense content and a flat OAM close and parallel to the plasma membrane. Even though, these two membranes do not interact until the sperm is stimulated in the proximity of the mature oocyte to undergo the acrosome reaction. After stimulation with a calcium ionophore or progesterone [9, 10], 30–40% of the sperm show morphologically altered acrosomes. Strikingly, the morphology of the equatorial region of the acrosome is similar to that observed in non-stimulated cells, meaning that this region remains intact. Some acrosomes undergo a simple swelling; however, in many cells, the OAM shows a wavy surface. The protrusions of this membrane occasionally come in contact with the plasma membrane and result in invaginations that are very deep (Figs. 1 and 5). Some cells appear to have vesicles inside the acrosome. Finally, a small group of cells show vesiculated or lost acrosomes, this state is known as reacted. Therefore, in response to exocytosis inducers, a granule first swells to get into contact with the cell plasma membrane, second, it gets attached to the plasma membrane and fuses with it. Finally, it detaches entirely, together with the portion of the plasma membrane that surrounds it. These events are coupled to a complex calcium signaling [11, 12]. In most secretory processes, fusion pore opening and expansion lead to the release of granule contents and the incorporation of the acrosomal membrane into the plasma membrane. In contrast, in mammalian sperm, pore opening and expansion cause the vesiculation of the acrosome. Acrosomal swelling and OAM deformation are important to delineate the membrane domains where pore expansion will lead to the release of hybrid vesicles (Figs. 1 and 5). Reproductive biologists have not yet reached a consensus about crucial issues including this remarkable deformation of the acrosomal granule occurring during the acrosome reaction, but it is thought to be related to lipid remodeling. 1.3 Kinetics of the Acrosome Granule Exocytosis

Most of the knowledge of exocytosis comes from scientific reports on secretion in neurons and neuroendocrine cells. These cells contain several small vesicles that fuse quickly with the plasma membrane. Much less is known about secretion of the large granules stored in exocrine cells, such as pneumocytes and acinar and salivary cells. There is an important correlation between granule size and secretion kinetics [13]. The acrosome reaction is not an exception to this rule: large acrosomal granules exhibit slow (minutes)

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kinetics of release [10]. Results from several laboratories have shown that the percentage of reacted sperm increases with the time of incubation in the presence of physiological and pharmacological stimuli [14–16]. The kinetics of the process is measured in minutes and depending on the experimental conditions, it may require about 1 h to reach a plateau. Sosa et al. [10] measured the kinetics of human sperm exocytosis using different approaches. The swelling of the acrosomal granule, which precedes exocytosis, was a slow process (t½ ¼ 13 min). When the swelling was completed, the fusion pore opening occurred rapidly (t½ ¼ 0.2 min). Therefore, the acrosomal swelling is the slowest step and it determines the kinetics of acrosome reaction. After swelling is completed, the efflux of calcium from intracellular stores triggers fusion pore opening and the release of hybrid vesicles in seconds. 1.4 Why Is This Chapter in This Book?

Exocytosis of the acrosomal vesicle is unique, owing to the size and morphology of the granule, as well as the nature of the acrosomal contents; consequently, its exocytosis is a slow event. Despite the singular morphological and functional features, all of the fusion-related molecules involved in the acrosome reaction were initially found in somatic cells and implicated in various membrane fusion events. Therefore, sperm share their basic fusion machinery and regulatory components with all other eukaryotic cells, but their differences have to be highlighted [17–22]. Different cell types carry different secretory granules, such as neuronal, neuroendocrine, endocrine, and exocrine cells. Regulatory exocytosis varies among these cells in terms of cargo, kinetics, probability, and modality of release [12]. To release their contents, secretory vesicles must travel from a cytosolic compartment toward the plasma membrane. Because of its size and shape, the acrosome cannot travel to contact the plasma membrane. The acrosome reaction shares many features with exocytosis in other cell types, however, it differs at the following points: 1. Sperm contains only one secretory vesicle that overlies the anterior half of the nucleus in the apical region of the head instead of many secretory granules; 2. This granule has a unique secretion mode that differs from full collapse, kiss, and run, or compound exocytosis. On the contrary, the sperm acrosome reaction involves fusion at multiple points of the OAM and the overlying plasma membrane. Thus, many fusion pores are formed instead of one as in secretory somatic cells; 3. In contrast to full-collapse exocytosis where the release of granular contents into the extracellular medium causes the incorporation of their membranes into the plasma membrane, acrosome reaction is characterized by the release of hybrid

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vesicles (half OAM and half plasma membrane) into the media with membrane loss. 4. Acrosome reaction is a tightly regulated and irreversible process with unknown membrane recycling. Once the granule releases the acrosomal contents, the cell modifies the membrane components entirely, exposing the IAM to the extracellular medium and keeping the equatorial segment intact (Fig. 1). The end point of most studies involving acrosome reaction is the staining of cells at different time points after stimulation to calculate the percentage of reacted sperm. This experimental setting measures three time components: (1) the time required for a stimulated sperm to activate the molecular machinery of membrane fusion, (2) the time required for the actual opening of fusion pores, and (3) the time required for the expansion of fusion pores to release the acrosomal contents. It also includes the probability of each cell to respond to the stimulus received. It is essential to clarify that not all human sperm react at the same time, but in fact a small percentage (10–30%) undergo the acrosome reaction [10, 16, 23]. The reason for this asynchrony is not understood. Spermatozoa are terminal cells lacking almost all organelles, transcriptional and translational activities. From the point of view of intracellular trafficking, sperm are specialized for a single membrane fusion event, the exocytosis of the acrosome granule. This feature is particularly useful to study exocytosis in isolation. At the same time, their inability to synthesize proteins constitutes the biggest obstacle when trying to apply classic cell biology and biochemistry approaches to sperm. Our laboratory has successfully contributed to establishing two techniques to overcome this limitation: controlled plasma membrane permeabilization with pore-forming toxins, like streptolysin or perfringolysin O, and delivery of permeable proteins [17, 24–30]. The latter allows sophisticated molecular studies of the pathways elicited by well-established AR inducers because it grants access to intracellular compartments in non-permeabilized cells. Given the particular characteristics of sperm exocytosis discussed above, methods commonly used to measure secretion in other cells cannot be used for this purpose. Here, we provide detailed protocols for studying human spermregulated exocytosis at a defined time or at real time by using different approaches.

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Materials Ultrapure water must be used to dilute all reagents. Prepare it by purifying deionized water to attain a sensitivity of 18 M Ω cm at 25  C. Use analytical-grade reagents. Follow all waste disposal regulations when discarding materials.

2.1 Human Tubal Fluid (HTF Media)

For 1 L of HTF weigh 5.94 g/L NaCl, 0.35 g/L KCl, 0.05 g/L MgSO4·7H2O, 0.05 g/L KH2PO4, 0.3 g/L CaCl2·2H2O, 2.1 g/ L NaHCO3, 0.51 g/L D-glucose, 0.036 g/L Na pyruvate, 2.39 g/ L Na lactate, 0.06 g/L penicillin, 0.05 g/L streptomycin, and 0.01 g/L phenol red (see Notes 1–3).

2.2 Hepes BufferEthylene Glycol-bis(2aminoethylether)-N,N, N 0 ,N 0 -tetraacetic Acid (HB-EGTA)

250 mM sucrose, 20 mM HEPES free acid, 0.5 mM EGTA, pH 7. Dissolve all the reagents in water at room temperature by stirring (see Note 4). Adjust pH with 1 N KOH.

2.3 PhosphateBuffered Saline

For 1 L of PBS (1) weigh 8 g NaCl, 0.2 g KCl, 1.44 g Na2HPO4 and 0.24 g KH2PO4. Adjust pH to 7.4 with HCl. Finally, add distilled water to a total volume of 1 L.

2.4 Exocytosis Inducers Used for Non-permeabilized Spermatozoa (See Note 5)

1. A23187 (see Note 6). For 100 mM stock solution 1: 1 μL of 100 mM stock solution 1–9 μL of dimethyl sulfoxide (DMSO) to yield a 10 mM solution (stock solution 2). Tip off 2.5 μL of the stock solution 2 and pour it on 47.5 μL of HTF without bovine serum albumin (BSA) to yield a final concentration of 0.5 mM (Stock solution 3). Add 1 μL of the stock solution 3 to the experimental condition tube containing 49 μL of sperm suspension to obtain a final concentration of 10 μM. Stock solutions can be stored up to 3 months at 20  C (see Note 7). 2. Progesterone (Pg, see Note 8). Prepare 10 mM stock solution in absolute ethanol and gently swirl to dissolve. Tip off 2.5 μL of the stock solution and pour it on 47.5 μL of HTF without albumin to yield a final concentration of 0.5 mM. Add 1.5 μL of 0.5 mM Pg solution in 48.5 μL of sperm suspension in HTF with BSA to obtain a final concentration of 15 μM. Store aliquots at 20  C. Avoid repeated freeze–thaw. 3. Diacylglycerol (DAG, see Note 9). Prepare a 0.5 mM solution in HTF without BSA (dissolve 1 μL of a 25 mM DMSO stock solution in 49 μL of HTF without BSA). To obtain a final concentration of 10 μM DAG in the experimental tube, add

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1 μL of the 0.5 mM solution in HTF to 49 μL of sperm suspension. 4. Phorbol 12-Myristate 13-Acetate (PMA, see Note 10). Prepare a 1 M stock in DMSO, apply a N2 stream, and store at 20  C. Add 0.5–49.5 μL of DMSO to obtain a 0.01 M solution. From this stock, prepare additional dilutions in DMSO until a 10 μM PMA solution is obtained. Add 1 μL from the 10 μM stock to 49 μL of sperm suspension to get a final concentration of 200 nM. 5. Sphingosine-1-Phosphate (S1P). Prepare a solution by adding 995 μL of methanol and 5 μL of water and add it to the vial containing sphingosine-1-phosphate (S1P) (1 mg), provided by the manufacturer. Heat at 50  C in a water bath, vortex, and then sonicate in a water bath sonicator to dissolve the contents. Label ten glass vials covered with aluminum foils or use amber glass vials. Aliquot 100 μL into the glass vials. Dry to powder under a stream of N2 gas. Seal vials. Freeze and store at 80 ºC. Resuspend dried aliquots in 100 μL HTF supplemented with 0.5% fatty acid free BSA to yield 2.6 mM S1P stock solution [31]. 6. Soluble Human Recombinant Zona Pellucida (ZP, see Note 11). We have performed functional experiments to assess ZP’s ability to induce acrosomal exocytosis for isoforms 2, 3, and 4 (unpublished data). To perform functional assays, we incubate 1  106sperm with 1 μg/μL final concentration of ZP2 (negative control), ZP3, or ZP4. 2.5 Exocytosis Inducers Used for Permeabilized Spermatozoa (See Note 12)

1. Calcium. Dissolve 2.77 mg CaCl2 in 1 mL ultrapure water to obtain a 25 mM stock solution. Tip off 1 μL and pour it on 49 μL of sperm cells suspension in HB-EGTA to yield a final concentration of 0.5 mM (see Note 13). 2. Diacylglycerol. Prepare this solution as described under Subheading 2.4, item 3 (for non-permeabilized cells), an exception is the final stock solution which must be prepared in HB-EGTA. 3. Phorbol 12-Myristate 13-Acetate. Prepare this solution as described under Subheading 2.4, item 4. The final stock solution must be prepared in HB-EGTA. For further information see Note 14.

2.6 FluoresceinIsothiocyanateCoupled Pisum Sativum Agglutinin

Prepare a 5 mg/mL PSA stock solution in PBS and store aliquots at 20  C. PSA working solution: Dilute 1 μL of PSA stock solution in 199 μL of PBS (intact sperm) or HB-EGTA (permeabilized sperm). This dilution must be used directly to stain the samples. Wrap the tubes with aluminum foil to protect from light.

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2.7 Coomassie Blue G250 Solution

Weigh 0.11 g Coomassie Blue G 250 and dissolve in methanol/ glacial acetic acid/distilled water (v/v 25:5:20 mL).

2.8 Four Percent Paraformaldehyde Solution

Weigh 4 g PAF and 0.5 g NaOH. Add 90 mL deionized water. Transfer to an Erlenmeyer flask and cover it with aluminum paper. Heat the solution on a magnetic stirrer under fume hood until the solution is transparent (60  C). Adjust pH to 7. Add 10 mL 1 PBS to reach 100 mL as final volume. Freeze aliquots.

2.9 Ammonium Acetate (0.1 M)

Weigh 0.07 g ammonium acetate (CH3CO2NH4) and add 10 mL of deionized water.

2.10 Sodium Cacodylate Solution

Weigh 21.40 g sodium cacodylate ((CH3)2AsO2Na·3H2O) and add 1 L of deionized water to reach 0.1 M final concentration, pH 7.4.

2.11 Glutaraldehyde Solution (2.5%)

Use electron microscopy grade 25% glutaraldehyde in sealed ampoules. Add 1 mL of 25% glutaraldehyde to a solution of 9 mL sodium cacodylate (0.1 M, pH 7.4).

2.12 Osmium Tetroxide Aqueous Stock Solution

Work under a fume hood and use nitrile gloves. Open a sealed glass ampoule of 0.5 g OsO4 and add it to deionized water. Complete with water to reach a final volume of 25 mL (2% final concentration).

2.13 Uranyl Acetate Aqueous Solution

Weigh 1 g uranyl acetate and add to 50 mL of ultrapure water by stirring to reach a final concentration of 2%.

2.14 Lead Citrate Solution

Add 0.1 g NaOH and 0.18 g lead citrate to 25 mL of distilled and boiled water.

3

Methods

3.1 Sample Collection

1. Human semen samples should be obtained by masturbation and ejaculated into a clean, wide-mouthed container made out of glass or plastic, from a batch that has been confirmed to be non-toxic for spermatozoa (see Note 15). 2. Label the specimen container with an identification number, the date, and time of collection. 3. Place the specimen container in an incubator (37  C, 5% CO2) to liquefy the semen (30 min to 1 h) (see Note 16). 4. After liquefaction, evaluation of semen quality must be done (see Note 17).

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3.2 Swim-Up (See Note 18)

1. Mix the semen sample well (see Note 19). 2. Place 1 mL of semen in a 5-mL polypropylene tube (12  75 mm). Alternatively, a conical centrifuge tube can be used. 3. Gently layer 1.2 mL of BSA-supplemented medium over the semen sample. Alternatively, pipette the semen carefully into the BSA-supplemented culture medium. 4. Place the tubes at 45 angle, to increase the surface area of the semen-culture medium interface, and incubate for 1 h at 37  C. 5. Return the tube to the upright position gently and remove the uppermost 1 mL of medium. This will contain highly motile sperm cells. 6. Mix the recovered cells well for assessment of sperm concentration (see Notes 20 and 21). 7. The specimen may be used directly or after capacitation for research purposes.

3.3 Capacitation of Spermatozoa

For acrosome reaction assays, spermatozoa must be capacitated (see Note 22). 1. Prepare the HTF-BSA capacitation-inducing medium fresh for each assay (see Note 3). 2. Warm up the medium to 37  C before use, preferably in a 5% (v/v) CO2-in-air incubator. 3. Dilute the motile sperm population obtained from swim-up to 10  106 cells/mL using fresh warm HTF-BSA medium. 4. Incubate the sperm suspensions for at least 3 h at 37  C in an atmosphere of 5% (v/v) CO2-in-air incubator to induce capacitation (see Note 23) [32].

3.4 Functional Assays

Once capacitation time has ended, cells can be treated with inhibitors or stimulants of acrosomal exocytosis immediately, or permeabilized. In both cases, cells can be treated as follows: 1. Prepare experimental tubes, each containing the volume established for each reagent from stock solutions (stimulants or inhibitors of different components of the signal transduction cascade, see Subheading 2.4). Identify different conditions with numbers . 2. Add to each tube the volume of sperm suspension required to reach 50 μL (7–10  106/mL motile spermatozoa, 350,000 to 500,000 cell/condition). 3. Mix gently with a pipette changing the tip for each condition. 4. Incubate all the tubes for 15 min at 37  C, 5% CO2/95% air.

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5. If the cells are treated with an inhibitor, subsequently add the microliters stated in Subheading 2.4 for the stimulus to be used and incubate for an additional 15 min at 37  C, 5% CO2/ 95% air. 6. For control conditions, incubate the sperm without the addition of any reagent (see Note 24). 7. After that, 10 μL of each reaction mixture must be spotted on 8- or 12-well slides (microscope slides, coated with Teflon, size: 75  25  1 mm). 8. Air-dry the spots. 9. When dried, fix/permeabilize the cells covering the slide with 1 mL of ice-cold methanol for 1 min. 10. Wash four times  5 min each with distilled water by placing 1.5 mL on the slide after turning over the slide to eliminate the water of the previous wash (see Note 25). 11. Using a conventional transillumination microscope check if the preparation is clean enough. 3.5 Permeabilization of Spermatozoa (See Note 26)

1. Recombinant streptolysin O (SLO) was obtained from Dr. Bhakdi (University of Mainz, Mainz, Germany). 2. Wash the sperm suspension 2 with ice-cold PBS (use the spermatozoa recovered from swim-up; capacitate and dilute to 7–10  106, as described in Subheadings 3.2 and 3.3. 3. Suspend the pellet by gently pipetting in a volume of cold PBS equal to the original one. 4. Before using a new batch of the protein performe a doseresponse curve to determine the optimal SLO concentration to be used. 5. Add 0.7 μL of pre-activated SLO (300 U/mL) per 100 μL of sperm suspension to get a final concentration of 2.1 U/mL (see Note 27). 6. Incubate for 15 min at 4  C. 7. Remove the remaining non-bound toxin from the cell suspension by washing once with cold PBS. 8. Resuspend the pellet in ice-cold HB-EGTA (see Subheading 2.2) containing 2 mM dithiothreitol (DTT) to provide a reducing environment. 9. Label the tubes with numbers and add to each tube the volume of stimulants or inhibitors as described in Subheading 2.5. 10. Aliquot sperm suspension (prepared in HB-EGTA plus 2 mM DTT) in the experimental tubes containing reagents to a final volume of 50 μL. First add an inhibitor and incubate with the cells for 15 min at 37  C, 5% CO2/95% air. After that, the stimulus must be added and tubes incubated for an additional 15 min under the same conditions.

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11. Include in all experiments negative (no stimulation) and positive controls (stimulated with 0.5 mM CaCl2 rendering 10 μM free calcium (see Subheading 2.5, item 1)). 12. Finally, 10 μL of each reaction mixture must be spotted on 8or 12-well slides (microscope slides, coated with Teflon, size: 75  25  1 mm). 13. Let the spots air-dry. 14. When dried, fix/permeabilize the cells covering the slide with 1 mL of ice-cold methanol for 1 min. 15. After that, follow the protocol for assessing the acrosomal exocytosis as described in Subheading 3.4, steps 7–11. 3.5.1 SLO Pre-activation

1. Add 1 volume of 6 mM DTT to 2 volumes of 450 U/mL SLO, obtaining a final SLO concentration of 300 U/mL. 2. Incubate the mixture at 37  C for 30 min. After that incubation period, store the protein on ice until use.

3.5.2 SLO Dose– Response Curve (See Note 28)

1. Before using a new batch of the protein, a dose–response curve must be generated. Generate a curve from 0.1 U/mL to 10 U/ mL. 2. Evaluate the percentage of permeabilized cells by eosin yellowish staining (see Note 29). 3. Choose the SLO dilution that renders 70–80% of permeabilized cells (meaning the percentage of cells where eosin has entered via membrane pores). 4. Perform a functional assay as described in Subheading 3.4 using calcium as a stimulus. 5. Aliquots of concentrated stock of SLO (e.g., 25,000 U/mL) must be stored at 80  C. Avoid repeated freeze–thaw (see Note 30).

3.6 Assessment of the Acrosome Reaction (See Note 31)

Follow this protocol for the cells obtained and treated as described in step 6 of Subheading 3.4.

3.6.1 Visualization of Acrosomal Status Using Light Microscopy. Coomassie Blue Staining Method (See Note 32)

2. Wash the pellet with HTF (RT) without BSA and resuspend it in the same media (10  106 cells/mL).

1. Centrifuge the sperm suspension at 700  g for 3 min.

3. Prepare experimental tubes by adding the volume of sperm suspension required to reach 1  106 motile spermatozoa/ mL per condition (100 μL). 4. Add the volume established for each inducer from stock solutions (stimulants, see Subheading 2.4). Identify different conditions with numbers. 5. Mix gently with a pipette, changing the tip for each condition.

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6. Incubate all the tubes for 15 min at 37  C, 5% CO2/95% air. 7. Centrifuge sperm suspension at 2600  g for 3 min. Wash the pellet with HTF without BSA. 8. Suspend the pellet in 4% PAF (see Subheading 2.8) (100 μL). Incubate the sperm for 1 h at 4  C. 9. Centrifuge at 1700  g for 2 min. 10. Wash the cells once first with PBS 1 and then twice with 0.1 M ammonium acetate pH 9. 11. Centrifuge the cells at 700  g for 3 min. 12. Prepare sperm smears with 100 μL of suspension on poly-Llysine-coated microscope slides (see Note 33). 13. Allow the slides to air-dry. 14. Inspect the smears under a phase-contrast microscope (400). 15. Ensure that the spermatozoa are evenly distributed on the slides without clumping. 16. Wash the slides with distilled water for 5 min. 17. Incubate with methanol for 5 min. 18. Wash with distilled water for 5 min. 19. Let the slides air-dry and then cover them with a Coomassie blue solution (see Subheading 2.7) for 10 min. 20. Wash the excess of Coomassie blue by immersing the slides in distilled water at RT. For this purpose, use a glass coplin jar. 21. Let the slides air-dry. 22. Drop Mowiol (prepared as the manufacturer instructions) on the slide, place over a coverslip, and observe under light optical microscope. 23. Scoring (see Note 34 and Fig. 2.) 3.6.2 FluoresceinIsothiocyanate-Coupled Pisum Sativum Agglutinin Assessment of Acrosomal Status by Indirect Staining Method Using Epifluorescence Microscopy (See Notes 35 and 36)

Follow this protocol for the cells obtained and treated as described in Subheadings 3.4 or 3.5. 1. Prepare a 50 μg/mL solution of FITC-PSA in 1 PBS (see Subheading 2.6) see Note 35. 2. Pour 10 μL of the lectin dilution solution on each spot. 3. Incubate in a moist chamber in the dark for 40 min at RT. 4. Wash the excess of lectin by immersing the slide in distilled water for 20 min at 4  C. For this purpose, use a glass coplin jar with lid covered with aluminum foil to keep in dark. 5. Allow drying in a dark chamber. 6. View the slide using a microscope equipped with epifluorescence optics at 600 magnification with oil immersion at 450–490 nm excitation.

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Fig. 2 The micrograph shows that Coomassie Blue G-250-stained acrosomeintact sperm are easily discernible from acrosome-reacted sperm. Human sperm were washed, fixed, and stained with the dye. Arrows indicate acrosome-intact sperm. Asterisk indicates acrosome-reacted sperm that shows negligible staining in the acrosomal region as compared to the acrosome-intact sperm

7. Categorize the spermatozoa as described in Note 36. 8. Tally the number in each acrosomal category (AI and AR) with the aid of a laboratory counter. 9. Score at least 300 cells in order to achieve an acceptably low sampling error. Include in all experiments negative (no stimulation) and positive controls (stimulated with 10 μM A23187 or 15 μM Pg). For permeabilized sperm, use calcium as positive control as indicated in Subheading 2.5, item 1 (see Note 37). 10. For each experiment, calculate acrosomal exocytosis indexes as described in Note 38. 3.6.3 FluoresceinIsothiocyanate-Coupled Pisum Sativum Agglutinin Assessment of Acrosomal Status by Indirect Staining Method Using Flow Cytometry

1. Cells can be stained by using the same technique in suspension as described in Subheading 3.6.2 and scored by flow cytometry. 2. Wash the sperm suspension once with warm HTF (no BSA addition) for intact sperm. In case of permeabilized cells, resuspend them in HB-EGTA. Use 100 μL/per condition of spermatozoa recovered from swim-up, capacitated and diluted to 7–10  106 as described in Subheadings 3.2 and 3.3. We recommend creating multiple aliquots of different cell conditions in order to obtain measurements in duplicate. 3. Follow this protocol from cells obtained and treated as described in Subheadings 3.4 or 3.5. 4. Centrifuge at 1700  g for 3 min.

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Fig. 3 Micrograph of human spermatozoa showing indirect FITC-PSA staining patterns. The lectin stains the intact acrosome (arrows). The micrograph also shows spermatozoa that have lost the acrosome (reacted, asterisk)

5. Fix/permeabilize with 100 μL of cold methanol for 1 min. 6. Centrifuge at 1700  g for 3 min. 7. Wash the cells 1–3 times by resuspending them in 300 μL of distilled water per tube and mix them gently. Centrifuge the samples at 700  g for 3 min and aspirate the supernatant. 8. Prepare a 50 μg/mL solution of FITC-coupled PSA in 1 PBS (see Subheading 2.6). 9. Add 50 μL of the lectin dilution solution to each tube. Resuspend by pipetting gently. 10. Incubate in the dark for 40 min at RT. 11. Centrifuge at 700  g for 3 min and discard the supernatant. 12. Wash the excess lectin 2 by adding 500 μL of distilled water and then centrifuge at 700  g for 3 min. Discard the supernatant. Protect the tubes from light. 13. Suspend in 200 μL PBS 1 and run each sample condition in flow cytometer. 14. As an auto fluorescence control, one aliquot of sperm must be left with no lectin addition. Once capacitation time has ended, cells can be treated with inhibitors or stimulants of acrosomal exocytosis immediately.

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3.6.4 FluoresceinIsothiocyanate-Coupled Pisum Sativum Agglutinin Assessment of Acrosomal Status by Direct Staining Method Using Epifluorescence Microscopy and Flow Cytometry (See Note 39)

1. Wash the sperm suspension 1 with warm HTF (no BSA addition) for intact sperm. In case of permeabilized cells, resuspend them in HB-EGTA. Add 5 μg/mL of FITC-PSA. Use 100 μL/per condition of spermatozoa recovered from swimup, capacitated and diluted to 7–10  106, as described in Subheadings 3.2 and 3.3. 2. As an auto fluorescence control, one aliquot of sperm should be left with no FITC-PSA addition. 3. Other aliquots: incubate with inhibitors for 15 min at 37  C. Protect them from light by covering the tubes with aluminum foil. 4. Incubate with stimulants for additional 15 min at 37  C. 5. Wash twice each condition with either HTF or HB-EGTA. 6. Spin down the cells at 700  g for 3 min. 7. Resuspend pellets with 2% PAF. 8. Incubate for 15 min at 4  C. 9. Spin down for 3 min at 3300  g. 10. Resuspend pellets in 200 μL 1 PBS (see Note 40). 11. Analyze conditions (10.000 events per condition) in a flow cytometer (we analyze the data by FlowJo software). Keep the cells at 4  C protected from light until they are scored in the cytometer (see Note 41). 12. It is very important to vortex each sample before running in the cytometer. 13. The gating scheme must be set up in the instrument prior to data acquisition. First, run the control condition (no PSA addition) to gate the cell population and discard cellular debris. 14. Then the rest of the conditions can be run according to the experiment. 15. Dot plot graphs of a representative experiment are shown in Fig. 4. To assess exocytosis by epifluorescence microscopy, cells obtained at step 10 of this protocol can be spotted on 8- or 12-well slides. Check them under the epifluorescence microscope to evaluate the staining pattern. If the fluorescence is faint, stain again as indicated in Subheading 3.6.2. In the direct method, fluorescent acrosomes are considered reacted and no fluorescent acrosomes are scored as intact.

3.6.5 Real-Time Measurements of the Acrosome Reaction (See Note 42)

1. Immobilize the capacitated sperm (5  106 cells/mL) on polyL-lysine (0.01% w/v) precoated cover slides (see Note 33). 2. Mount the samples in a chamber at 37  C and overlay them with HTF-BSA medium supplemented with 5 mg/mL FITCPSA (see Note 43).

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Q1 77,6

Q2 22,4

b 100K

80K

80K

60K

60K

SSC-A

SSC-A

a 100K

40K

40K

20K

20K

0

–103

0

103 FITC-A

104

105

0

Q1 63,8

–103

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Q2 36,2

0

103 104 FITC-A

105

Fig. 4 Measurement of the AR by a direct method using flow cytometry. Dot plot graphs of a representative experiment showing the number of FITC-PSA stained cells. Ten thousand cells were counted on each event and are represented by a single dot (side scatter (SSC) vs. FITC-PSA fluorescence). Note the low fluorescence index in control population (a) without any stimulus compared to the increased fluorescence that appeared upon incubation with a stimulus (b)

3. Perform image analysis by using the software Image J (National Institutes of Health, http://rsb.info.nih.gov/ij/) see Note 44. 3.6.6 Assessment of Acrosome Reaction and Ultrastructure of Sperm Using Transmission Electron Microscopy (See Note 45)

This protocol can be performed from the cells obtained and treated as described in Subheadings 3.4 or 3.5 as stated in Zanetti and Mayorga [9]. 1. After washing with 1 PBS, centrifuge the samples at 1700  g for 2 min to obtain at least a 20–25 μL of pellet. 2. For fixation, suspend the sperm pellet in 200 μL of 2.5% glutaraldehyde in sodium cacodylate (see Subheading 2.11). Incubate for 1 h to ON at 10  C. 3. Wash fixed sperm twice by suspending in sodium cacodylate for 15 min at 10  C (see Subheading 2.10). 4. Fix the samples with 1% osmium tetroxide in sodium cacodylate for 2 h, at RT under a fume hood (see Subheading 2.12). 5. Wash the samples three times by suspending in sodium cacodylate for 15 min at 10  C and spin down in a microfuge. 6. Dehydrate the pellets with increasing concentrations of acetone (50%, 70%, 95%, 100% and 100%), for 15 min in each step. 7. Embed the pellet with an epoxy resin (preferentially with low viscosity) gradually, with a change in resin/acetone (1:1 v/v) for 2 h. Then add pure resin and let embed ON at RT. 8. Polymerize in an oven at 70  C for 48 h to obtain a hard block.

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Fig. 5 Electron micrographs show the ultrastructural morphological correlate represented in the cartoons in Fig. 1. (a) Two cells classified as intact (unreacted) sperm (arrowheads). Both sperm heads show intact acrosomes (a), nucleus (n), and equatorial segment (es). (b) Swollen acrosome after stimulation (as). Invaginations of the outer acrosomal membrane contribute to the formation of the limiting surface of future hybrid vesicles (hv). Notice the close membrane apposition (ma) between the plasma and the outer acrosomal membranes at the edge of the invaginations. Some vesicles inside the acrosome (asterisk). (c) Micrograph showing sperm with a vesiculated acrosome. The membranes at the edge of invaginations detached from the head, and hybrid vesicles (hv) are released. (d) A sperm that has completed the acrosome reaction. The equatorial segment (es) remains unaltered during the vesiculation process (es)

9. Trim the block and section with an ultramicrotome. Collect the grey interference color sections (60-nm thickness) on 300 mesh copper grids. 10. Contrast the sections mounted on the grids with a solution of uranyl acetate (see Subheadings 2.13 and 2.14) for 5 min and then with a solution of lead citrate for 1 min. 11. Now, grids are ready to be observed at TEM. 12. Count at least 100 cells per condition. Acrosomal patterns should be scored and classified as intact, reacted (vesiculated and lost), and swollen (enlarged and deformed) (Fig. 5).

4

Notes 1. Use a glass beaker and add the reagents one at a time in water at RT while the magnetic stir bar is working. Make sure that each reagent is completely dissolved before adding the next one.

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HTF must be sterilized using aseptic processing techniques (filtration). Once prepared, the aliquots can be stored (15 mL) at 2–8  C until further use. Do not freeze the complete HTF or expose to temperatures greater than 39  C. To freeze and use aliquots later, combine all but NaHCO3 and freeze at 70  C. For experiment, thaw an aliquot and add appropriate NaHCO3. Let sit in 5% CO2 at 37  C ON. 2. The medium uses NaHCO3 as a buffering system. This is specifically designed for use in a CO2 incubator. The bottle/ tube of HTF medium should be loosely capped to allow for the exchange of gas and pH equilibration. Incubate HTF in 5% CO2 overnight at 37  C. Check to ensure the pH is between 7.4 to 7.6. 3. Human Tubal Fluid media is used to perform the swim-up technique and to isolate motile fraction of a semen sample. To perform acrosome reaction assays, the sample needs to be capacitated, which is achieved in vitro by incubation of the motile fraction recovered with HTF supplemented with 5 mg/mL bovine serum albumin (BSA) for at least 3 h [32]. 4. A 1 M HEPES-free acid solution (954 mg in 4 mL of water) is clear and colorless, with pH approximately between 5.0 and 6.5 at 20  C. Adjust pH with 1 N KOH. 5. The reagents we currently use to induce exocytosis in intact cells are able to penetrate the cell by different mechanisms; they are either permeant, able to open pores in the plasma membrane, or affect the function of some membrane surface proteins, for example, sphingosine 1-phosphate (S1P) receptors. Here, we provide a list of some reagents we usually use to analyze signal transduction cascades involved in exocytosis in human spermatozoa. 6. It is a calcium ionophore commonly used to induce exocytosis of the sperm granule. 7. DMSO concentration cannot be greater than 0.5% per experimental condition (it may produce cellular damage at higher concentrations). 8. Pg is the most widely used physiological inducer of the acrosome exocytosis. 9. DAG is an allosteric activator of protein kinase C (PKC) and induces the acrosomal exocytosis in intact and in permeabilized sperm [25, 31]. Storage: diacylglycerols are conveniently stored in chloroform solutions in glass vials with Teflon-lined caps at 20  C. Avoid plastic when handling chloroform solutions. Delivery to cells: dry DAG aliquots in chloroform, using a stream of N2. Dissolve the residue in an appropriate volume

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of DMSO, then dilute to the desired concentrations using aqueous medium. 10. PMA induces the acrosomal exocytosis. It has a structure analogous to DAG and can also activate PKC [27, 31]. PMA is soluble in DMSO. Prepare a stock solution and store in the freezer, protected from light. 11. Zona Pellucida. The zona pellucida are glycoproteins known as physiological inducers of the acrosomal exocytosis. In humans, there are four isoforms that are secreted by oocyte in order to build a specialized form of an extracellular matrix [33]. Human recombinant ZP proteins were obtained from Dr. Mayel Chirinos (Department of Reproductive Biology, Instituto Nacional de Ciencias Me´dicas y Nutricio´n Salvador Zubira´n, Mexico DF, Mexico). The protocol to obtain these proteins can be found in their publication [34]. The concentration of the single proteins to be used in non-permeabilized cells depends on each production batch. 12. We routinely use some recombinant permeant proteins involved in the signaling pathway leading to acrosome reaction, produced at our laboratory, as inducers or inhibitors of exocytosis. We cannot include the protocols to produce the proteins in this chapter, given that they exceed the topics to be developed here. If the readers are interested in that protocols, they can resort to the following bibliography [26, 30, 35]. 13. This dilution renders 10 μM free calcium, estimated by MAXCHELATOR, a series of programs for determining the free metal concentration in the presence of chelators available at http://maxchelator.stanford.edu. 14. The reagents used in permeabilized cells are different from those described in Subheading 2.4 Given that ion gradient is lost due to plasma membrane pores, these reagents become inactive. These reagents cannot be used as they are known to affect plasma membrane channels (because the plasma membrane is perforated). Further these inducers cannot be used as they interact with plasma membrane receptors. If an enzyme inhibitor is chosen for a defined signal transduction pathway, the component does not need to be permeant. Prepare the reagents following the manufacturer instructions and always generate a dose-response curve for the cells before use. Spermatozoa usually require inhibitors or stimulant concentrations that differ from other mammalian cells. 15. Semen samples were obtained from healthy donors with abstinence of at least 48 h. Data collection was carried out based on the principles outlined in the Declaration of Helsinki; all donors signed an informed consent agreeing to supply their own anonymous information and semen samples. The Ethics

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Committee of the Medical School, Universidad Nacional de Cuyo, approved the signed informed consent and the protocol for semen handling. 16. Occasionally, samples may not liquefy, making semen evaluation difficult. In these cases, mechanical mixing may be necessary. Inhomogeneity can be reduced by repeated (6–10 times) gentle passage through a blunt gauge 18 (internal diameter 0.84 mm) or gauge 19 (internal diameter 0.69 mm) needle attached to a syringe [36]. 17. Evaluation of semen quality is done as a routine practice in our laboratory. Detailed protocols to assess semen quality are described on WHO laboratory manual for the examination and processing of human semen [36] and are beyond the scope of this chapter. 18. Spermatozoa are selected by their ability to swim out of seminal plasma and into culture medium, meaning that we obtain the motile fraction. This is known as the “swim-up” technique. The direct swim-up technique is the technique of choice when the semen samples are considered to be largely normal. This is our case, for that reason is the procedure chosen instead of discontinuous density gradients or simply washing [36]. The semen should preferably not be diluted and centrifuged prior to swim-up. Motile spermatozoa then swim into the culture medium. 19. Before removing an aliquot of semen for assessment, mix the sample well in the original container, but not so vigorously that air bubbles are created. This can be achieved by aspirating the sample ten times into a wide-bore (approximately 1.5-mmdiameter) disposable plastic pipette. Do not mix with a vortex mixer at high speed as this will damage the spermatozoa. 20. Assessment of sperm concentration. We generally use the Makler counting chamber (is only 10 μm deep: one-tenth of the depth of ordinary hemocytometers, making it the shallowest of known chambers). It contains two pieces of optically flat glass, the upper layer serves as a cover glass, with a 1-sq.mm fine grid in the center subdivided into 100 squares of 0.1  0.1 mm each. Spacing is firmly secured by four quartz pins [37]. Counting Procedure: (a) Place a 10-μL drop of the immobilized, undiluted, and well-mixed preparation of liquefied semen at the center of the chamber by means of a pipette and cover it immediately. A microscopic objective of 20 is required. (b) Initiate the counting: the center of the chamber is subdivided into 100 squares of 0.1  0.1 mm each. Score sperm heads within ten squares in the same manner as blood cells are counted in a hemocytometer. The number

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obtained represents their concentration in millions per mL. (c) Rinse the chamber with water for reuse. Contact surfaces should be wiped with special lens paper after washing. (d) Alternatively, the Neubauer hemocytometer chamber can be used as described in the World Health Organization laboratory manual for the examination and processing of human semen [36]. 21. Makler chamber is as accurate as the Neubauer chamber; therefore, either one can be used in routine semen analyses [38]. We describe here the method we currently use at the laboratory. Even though the results obtained with both chambers do not differ significantly, [39] the Makler chamber has some practical advantages over the Neubauer chamber: (a) Applied spermatozoa are uniformly distributed and monolayered and are observed in one-focal plane. (b) Dilution is unnecessary even with concentrated specimens. Analysis is carried out directly from original specimen in its natural environment. (c) The specimen can be analyzed quickly even by an inexperienced person. 22. After leaving the testis, mammalian spermatozoa are morphologically differentiated but are immotile and unable to fertilize. Progressive motility is acquired during epididymal transit. However, freshly ejaculated mammalian sperm are not immediately capable of undergoing acrosome reaction and fertilizing an egg. They require a period of several hours in the female reproductive tract or in an appropriate medium in vitro to acquire this ability. The changes that occur during this period are collectively called capacitation [7]. This process is characterized by a special motility pattern and the ability to carry out the acrosome reaction under physiological stimuli [40, 41]. Many changes have been described during capacitation, including plasma membrane remodeling and flagellum hyperactivation. Major changes take place in the sperm plasma membrane, so as to render it fusogenic and responsive to ZP glycoproteins. However, the mechanisms involved have not been well defined. It is known to be a multistep process during which activation of a bicarbonate-dependent adenylyl cyclase leads to elevation of cAMP and protein kinase A (PKA)mediated tyrosine phosphorylation of a subset of flagellar proteins that correlates with changes in sperm motility. Upstream of these events, determinable changes occur to the sperm’s plasma membrane, like membrane hyperpolarization, opening of voltage-gated Ca2+ channels, loss of transbilayer

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phospholipid asymmetry, and cholesterol efflux (for a review see Visconti [7]). 23. Loosen the cap of the tube to allow gas exchange. If a CO2 incubator is not available, use a HEPES-buffered medium, cap the tubes tightly and incubate at 37  C. 24. Incubate the sperm with HTF alone to evaluate spontaneous acrosome reaction(background). To assess the maximum response, add only the stimulus (e.g., the calcium ionophore, A23187) and incubate for 15 min at 37  C. 25. This step is crucial because if the BSA present in the spot is not removed, it will get stained by FITC-PSA, generating a background, which will make it impossible to distinguish the sperm staining pattern. 26. Important advances in the field of exocytosis have depended on the use of permeabilized cells that allow the composition of the cytosol to be precisely controlled. A number of techniques have been devised to create pores in plasma membranes, such that cells become leaky, but do not lyse. Many cell functions are controlled by molecular signals (hormones, neurotransmitters, and so forth) that interact with cell-surface receptors and trigger specific intracellular responses. Intracellular signaling can be difficult to study in isolated, purified systems, because these events depend on cellular architecture to a large extent. In intact cells, access to intracellular systems is limited by the restricted permeability of the plasma membrane [42–45]. Given that the sperm is a transduction and translationally inactive cell and that some reagents, proteins, and lipids required to dissect exocytosis molecular mechanisms are not permeant, a controlled plasma membrane permeabilization is necessary. 27. The units of SLO indicated are calculated for the batch of the protein we are using nowadays. Calculate your own units for your batch. The incubation performed in this step allows SLO binding to cholesterol molecules present in the plasma membrane. 28. Streptolysin O belongs to the homologous group of thiolactivated toxins that are elaborated by various Gram-positive bacteria. This toxin binds as monomers to cholesterol in the cytoplasmic membranes. They then oligomerize into ringshaped structures, estimated to contain 50–80 subunits, which surround pores of approximately 30 nm diameter [46, 47]. 29. Mix 10 μL of sperm dilution with 2 μL of eosin yellowish 0.5% in PBS on a slide placed on a 37  C plate. After 1-min incubation, score cells in a conventional transillumination microscope with a 40 microscope objective.

If you do not want to detect spontaneous acrosome reaction occurring before applying the stimulus, you should use the direct method. It only detects sperm reacting during the incubation. Any sperm that had reacted before is not stained by the lectin present in the medium [23] Real-time measurements of exocytosis require the use of the direct method [27]

More expensive; due that a Fast, easy, and reliable fluorescence microscope Allows an accurate counting by flow equipment is needed or a flow cytometry, widely used cytometer Permits measurement of the exocytosis It is not used routinely in diagnostic in real time (kinetic studies) laboratories

Fluorescence assessment of acrosomal status. Pisum sativum Agglutinin coupled to fluorescein isothiocyanate (FITC-PSA) Direct and indirect staining

If you need to study the ultrastructure Acrosome reaction assessment by Allows the observation of the sequence It requires a week to be finished Experienced and trained professionals of the exocytosis, this is the method transmission-electron of events that occurs in a of choice microscopy spermatozoa’s head during human You can score and analyze different sperm acrosome reaction. changes of the granule like swelling, Particularly, the acrosome membrane curvature after different morphology at the ultrastructural treatments, number and depth of level invagination of the OAM, etc

Only to assess sperm exocytosis

Until now, the method has not been Fast, easy, reliable, and inexpensive set up for flow cytometry; that Requires only optical light microscope would allow an accurate counting Can be used in fertility and biochemical independent of the human eye laboratories for a routine semen analysis

Method of choice

Coomassie blue staining method

Disadvantage

Advantage

Method

Table 1 Comparison between acrosomal exocytosis assessment methods

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30. Considering the hypothetical stock of 25,000 U/mL a diluted stock can be prepared to work. A 450-U/mL stock must be prepared as follows: mix 9 μL of 25,000 U/mL SLO, 100 μL of glycerol, and 1 μL of 5% BSA in PBS. Add PBS to a final volume of 500 μL. Aliquot and store at 20  C if it can be used within a month; if not store at 80  C. 31. The read out of our functional assays is the measurement of the acrosomal status after different treatments. Knowing whether the acrosomes have been exocytosed or not after the incubation with diverse reagents allows building a signal transduction network. The acrosomal status can be evaluated by different methods each with advantages and disadvantages (Table 1). The appropriate method should be chosen depending on the requirement. The techniques developed below could be used both for intact and permeabilized cells. 32. The protocol was adapted in our laboratory from the technique published by Larson and Miller [48]. This method is fast, easy, and inexpensive. It does not require a fluorescence microscope; a light optical microscope will suffice. Therefore, this technique can be used in ordinary diagnosis laboratories of biochemistry together with a routine semen analysis. 33. Prepare 10 mL of poly-L-Lysine solution by adding 9.5 μL of distilled water to 500 μL of 0.1% (w/v) poly-L-Lysine. Smear this solution over ethanol-cleaned slides. Let them dry. 34. View the slide under a light optical microscope at 400 or 600 magnifications. Categorize the spermatozoa as follows. (a) Acrosome-Intact: spermatozoa in which more than half the head is uniformly blue-stained (see Fig. 2). (b) Acrosome-Reacted: spermatozoa with light blue staining in the acrosome region (see Fig. 2). (c) Abnormal Acrosomes: all other spermatozoa. Perform scoring and classify the cells in the categories described. 35. Pisum sativum agglutinin has four subunits, two of approximately 17,000 daltons and two of about 6000 daltons. Lectin has specificity toward terminal α-D-mannosyl-containing oligosaccharides present in the acrosome granule. If the acrosome is present, the FITC-coupled lectin binds to the mannose residues and the structure shows fluorescent green staining. On the contrary, if the acrosome is lost, mannose residues are not present, and the lectin does not stain the granule. 36. The method is generally used to evaluate the acrosomal status by staining the sperm cells with FITC-coupled PSA, according to Mendoza et al. [49]. It is fast, easy, and yields accurate information about sperm exocytosis. The laboratory must be equipped with a fluorescence microscope or a flow cytometer

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(Table 1). Two procedures can be used for this staining: indirect or direct. In the indirect staining method, we first perform the functional assay as described in Subheading 3.4. Then, we fix/permeabilize the sperm and finally, we stain them. The spermatozoa that preserve the acrosome even after applying the stimulus are considered non-reacted or intact acrosome (Fig. 3, arrows, fluorescent acrosomes). On the contrary, if the spermatozoa lose the acrosomes, they are considered as reacted (Fig. 3, asterisk, no fluorescent acrosome). In the direct method, the lectin that binds the mannose residues of the acrosome is present in the media during the functional assay. Therefore, when the acrosome starts the exocytosis, the lectin enters through the pores, staining the granules of reacted cells. For this reason, fluorescent acrosomes are considered reacted and no fluorescent acrosomes are scored as intact. The difference between the two methods is that the direct method only detects the sperm reacting during the incubation. Any sperm that had reacted before (spontaneous acrosome reaction) is not stained by the lectin present in the medium [23]. 37. Classification: a. Acrosome-Intact: spermatozoa in which more than half the head is brightly and uniformly fluorescent (Fig. 3). Acrosome-Reacted: spermatozoa with only a fluorescent band at the equatorial segment or no fluorescent stain at all in the acrosome region (Fig. 3). 38. Calculate acrosomal exocytosis indexes by subtracting the number of reacted spermatozoa in the negative control from all values and expressing the resulting values as a percentage of the acrosome reaction observed in the positive control. The average difference between positive and negative control must be around 15% or more (experiments where the difference between the negative and positive control is less than 10% must be discarded). 39. In the direct method, Pisum sativum agglutinin conjugated to FITC permeates into the acrosome when fusion pores open. The lectin stabilizes the acrosomal matrix preventing the dispersal of the granule contents [23]. 40. It is very important to remove the PAF completely (cells cannot be run onto the cytometer if they are resuspended in PAF). 41. If it is not possible to run the cells on the cytometer right away, they can be stored up to 48 h at 4  C, protected from light and resuspended in 1 PBS. 42. Another technique we would like to mention, given its importance, is the method published by Harper et al. [16] even though we do not use it as a routine practice. In this study,

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the progress of the AR is recorded in real time: fenestration of plasma membrane, exposure and dispersal of acrosomal content, and subsequent exposure of the IAM. A live human sperm is visualized using fluorescent labels for two different components of the acrosome. Soybean trypsin inhibitor (SBTI) binds to the acrosomal contents (specifically acrosin) of human sperm following membrane fenestration. The complement regulatory protein CD46 (membrane cofactor protein) is localized solely on the IAM in human sperm. As dispersal of the acrosomal content proceeds, binding sites for antiCD46 antibodies are revealed, allowing exposure of the IAM to be monitored. Simultaneous imaging of separate probes for acrosomal content and IAM show that rapid membrane fusion, initiated at the cell apex, is followed by exceptionally slow dispersal of acrosomal content (up to 12 min). 43. Induce the acrosome reaction with 10 μM A23187 or 15 μM Pg in intact cells or 10 μM calcium in permeabilized sperm (positive controls). Start to collect images once stimulus is added. We collect images in an Olympus FV1000 confocal microscope or an Eclipse TE300 Nikon microscope (2 frames/min) [27]. 44. Perform background subtraction by selecting the region of interest placed as close as possible to the sperm of interest. Discard any incompletely adhered sperm that moved during the course of any experiment. Carry out fluorescence measurements in individual sperm by manually drawing a region of interest around the head of each cell. When required, express raw intensity values as relative fluorescence normalized to the maximum fluorescence obtained after the A23187 or calcium addition. Record the time of acrosome reaction initiation for each cell and calculate a cumulative frequency (number of reacted sperm at time t/number of reacted sperm after 1 h  100). 45. For this technique, it is recommended to use a higher concentration of cells, at least 10  106 cells in 200 μL per condition. Refer to Subheading 3.4 and the same considerations must be taken for intact and permeabilized spermatozoa. Before processing the sample completely for TEM, perform a control of the functional assay by fluorescence microscopy as described in Subheadings 3.6.2 or 3.6.3. A complete revision of this technique can be found online http://www.bahiablanca-conicet. gob.ar/biblioteca/principios-practica-microscopia-electron ica.pdf

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SNAREs that mediate zymogen granule exocytosis. Biochem Biophys Res Commun 359 (3):599–603 43. Deeney JT, Branstrom R, Corkey BE, Larsson O, Berggren PO (2007) 3H-serotonin as a marker of oscillatory insulin secretion in clonal beta-cells (INS-1). FEBS Lett 581(21):4080–4084 44. Waselle L, Gerona RR, Vitale N, Martin TF, Bader MF, Regazzi R (2005) Role of phosphoinositide signaling in the control of insulin exocytosis. Mol Endocrinol 19 (12):3097–3106 45. Vitale N, Mukai H, Rouot B, Thierse D, Aunis D, Bader MF (1993) Exocytosis in chromaffin cells. Possible involvement of the heterotrimeric GTP-binding protein G(o). J Biol Chem 268(20):14715–14723 46. Sekiya K, Satoh R, Danbara H, Futaesaku Y (1993) A ring-shaped structure with a crown

formed by streptolysin O on the erythrocyte membrane. J Bacteriol 175(18):5953–5961 47. Sekiya K, Akagi T, Tatsuta K, Sakakura E, Hashikawa T, Abe A, Nagamune H (2007) Ultrastructural analysis of the membrane insertion of domain 3 of streptolysin O. Microbes Infect 9(11):1341–1350 48. Larson JL, Miller DJ (1999) Simple histochemical stain for acrosomes on sperm from several species. Mol Reprod Dev 52 (4):445–449. https://doi.org/10.1002/( SICI)1098-2795(199904)52:43.0.CO;2-6 49. Mendoza C, Carreras A, Moos J, Tesarik J (1992) Distinction between true acrosome reaction and degenerative acrosome loss by a one-step staining method using Pisum sativum agglutinin. J Reprod Fertil 95(3):755–763

Chapter 11 Bovine Chromaffin Cells: Culture and Fluorescence Assay for Secretion Tamou Thahouly, Emeline Tanguy, Juliette Raherindratsara, Marie-France Bader, Sylvette Chasserot-Golaz, Ste´phane Gasman, and Nicolas Vitale Abstract Over the last four decades, chromaffin cells originating from the adrenal medulla have been probably one of the most popular cell models to study neurosecretion at the molecular level. Accordingly, numerous seminal discoveries in the field, including the characterization of role of the cytoskeleton, fusogenic lipids, and soluble N-ethylmaleimide-sensitivefactor attachment protein receptor (SNARE) proteins, have been made using this model. In this chapter, we describe a standard method currently used to isolate and culture bovine chromaffin cells, and we illustrate a catecholamine secretion assay based on the successive transformation of adrenaline into adrenochrome and adrenolutine for fluorescence measurements. We also provide some guidelines for efficient cell recovery and for the use of this assay in the laboratory. Key words Chromaffin cells, Fluorescence assay, Catecholamine secretion

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Introduction Understanding how neurons communicate at the molecular level has been one of the most challenging goals of modern biology. A key step in improving this understanding was obtained by the discovery of some of the main players of the minimal protein machinery involved in synaptic vesicle and neuroendocrine granule exocytosis. Among the cell culture models that have provided insights into the molecular machinery underlying the successive steps of exocytosis, adrenal medulla chromaffin cells have taken a prominent place [1]. By offering the opportunity to combine the use of electrophysiological approaches with molecular biology for specific protein modifications and recent imaging techniques allowing single-vesicle resolution, chromaffin cells remain an influential model to address open questions in the field of secretion and exocytosis. In addition to the measurement of single-cell secretion

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by amperometry or capacitance, chromaffin cells have been widely used for cell population secretion assay. These assays were largely based on the use of either radiolabeled catecholamines or a reporter for secretion such as human growth hormone, specifically stored in secretory granules when expressed in chromaffin cells [2– 4]. Although the latter assay, by co-expressing the reporter with molecular tools, offers the possibility to measure secretory activity in a subpopulation of cells of interest, it remains important to be able to measure the secretory activity of a large population of non-transfected cells, for instance, to test the effect of pharmacological compounds on chromaffin cell secretion. Thus, to substitute for the use of radioactive compounds, we have developed a fluorescence assay based on the successive oxidation of noradrenaline and adrenaline into noradrenolutine and adrenolutine, respectively. In this chapter, we describe the detailed procedure for culturing bovine chromaffin cells and for measuring their secretory activity by a fluorescence-based method.

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Materials and Solutions Some procedures to culture rat and mouse chromaffin cells have been carefully described previously [5]. All procedures described here for bovine chromaffin cells require standard culture room facilities with a laminar flow cabin, a dry glass bead sterilizer burner, a water bath set at 37
C, and a 37
C incubator with a watersaturated atmosphere containing 5% CO2. An inverted microscope with phase contrast and two centrifuges are also required. Other required materials and basic solutions include: 1. Syringes and 0.22-μm syringe filters; 2. Adjustable volume pipettes with sterile tips; 3. 50-mL conical centrifuge tubes; 4. Hemocytometer (Neubauer); 5. Sterile surgical material required: forceps, scissors, and scalpel blades; 6. For the physical separation of cells, 250-mL centrifuge tubes are cut at the bottom; plugs are cut and used to adjust 217-μm mesh sieve filters. These ensembles are then autoclaved in aluminum foil; 7. 70-μm mesh filter; 8. 100-mm-diameter Petri dishes for dissection of glands; 9. A large glass tray (30 cm by 20 cm) and glass beakers (250 mL); 10. A centrifuge capable of reaching 20,000  g for Percoll gradient; 11. 96-well culture plates for cell culture.

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The following equipment is needed for the measurement of catecholamine levels in cultured chromaffin cells using a fluorescence-based assay: 1. 96-well black plates for the secretion assay; 2. Two Eppendorf Research Plus adjustable-volume multichannel pipettes and 5–10 multichannel disposable solution basins; 3. A fluorimeter allowing excitation at 430 nm and emission at 520 nm of 96-well plates. We use a LB 940 Mithras (Berthold) microplate reader. 2.1 Solutions and Medium for Culture and Secretion Assay

1. Ca2+- and Mg2+-free ten-fold concentrated Krebs solution: 1.54 M NaCl, 56 mM KCl, 36 mM NaHCO3, 56 mM glucose, and 50 mM HEPES (pH 7.4). This solution is filtered through 0.22-μm syringe filters. For each culture, 50 mL of concentrated Krebs solution is diluted in 450 mL of cell culture grade water to obtain 500 mL of Krebs balanced salt solution; 2. Trypan blue: 0.4% in Krebs balanced salt solution; 3. Cytosine arabinoside: 30 mg in 100 mL of Dulbecco’s modified Eagle medium (DMEM), and filter-sterilized; 4. Fluorodeoxyuridine: 24.6 mg in 10 mL of Krebs balanced salt solution and filter-sterilized; 5. 500 mL DMEM supplemented with L-glutamine plus 1 mL Primocin at 50 mg/mL to prevent Gram+ and Gram bacterial, mycoplasma, and fungal contaminations (DMEM-D). Depending on plating needs (see below), this medium is further completed with 1% cytosine arabinoside and 0.1% fluorodeoxyuridine solutions and 10% fetal bovine serum previously decomplemented for 30 min at 56
C to obtain DMEM-C. 6. 70 mg of collagenase A (0.235 U/mg) plus 100 mg of bovine serum albumin fraction V (BSA) diluted in 2 mL of concentrated Krebs solution. Adjust to 20 mL with Volvic (commercial) water, mix well, and filter-sterilize. Prepare 20 mL of collagenase solution per gland to be treated. 7. 18 mL of Percoll plus 2 mL of concentrated Krebs solution. Discard 1 mL and save 19 mL of the solution for later use. 8. Locke’s solution: 140 mM NaCl, 4.7 mM KCl, 2.5 mM CaCl2, 1.2 mM MgSO4, 1.2 mM KH2PO4, 0.01 mM EDTA, 11 mM glucose, 0.57 mM ascorbic acid, and 15 mM HEPES (pH ¼ 7.5). 9. Nicotine is diluted first in Locke’s solution at 1 M and stored protected from light. This intermediate dilution can be used within few hours to prepare the final 10 μM nicotine solution in Locke’s solution.

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10. High-potassium Locke’s solution: Prepare Locke’s solution with 59 mM KCl and NaCl adjusted to 85.7 mM to maintain osmolarity. 11. 1 M Sodium acetate solution adjusted to pH ¼ 6 with concentrated acetic acid. 12. 0.25% (w/v) of potassium ferricyanide (K3Fe(CN)6). 13. 5 M Sodium hydroxide containing 0.2% (w/v) of ascorbic acid. 14. Triton X-100 is diluted to 0.1% in water and stored at room temperature, protected from light. Bovine adrenal glands may be obtained from a local slaughterhouse and it is recommended that they be collected from the carcass immediately after the animal’s death. The shape of the two adrenal glands is different, with the left “bean-shaped” gland being more suitable for the following procedure. Collect only intact glands to avoid contamination and discard those with visible signs of internal blood coagulation. Leave significant level of fat at this stage as it provides protection against contamination. We recommend transporting the glands in an ice-cold plastic container with 200 mL of Krebs balanced salt solution containing Primocin if the delay between animal’s death and the start of the culture exceeds 1 h. Other protocols for bovine cell culture preparation have been published elsewhere [5–12]. Once in the culture room, remove the surrounding fat from the glands under a laminar flow cabin. The general procedure to minimize contamination is to proceed progressively from a nonsterile (transport containers, external fat) to a sterile environment. It is advisable to sterilize the surgical instruments frequently in the dry bead sterilizer. Once the glands are clean of surrounding fat and connective tissue, remove all the material used and discard the tissue from the hood.

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Methods

3.1 Bovine Chromaffin Cell Culture

The procedure described below is intended for six glands but can be easily adapted for more glands: 1. Prepare the collagenase solution. Each gland will require 20 mL of collagenase solution. These calculations are aimed for a commercial collagenase with an activity of 0.235 U/mg and should be adjusted according to enzyme activity (see Note 1); 2. Place the glands in a sterile Petri dish and inject each gland twice with 3–4 mL of Krebs balanced salt solution prewarmed at 37
C to remove blood cells;

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3. Inject the glands with 3–10 mL of collagenase solution depending on the size of the gland and place the glands standing in a 250-mL beaker. Try to avoid accumulation of liquid at the bottom of the beaker, which could contaminate the gland, and cover the beaker with a sterile aluminum foil; 4. Repeat step 3 after 10 min. Caution should be taken when performing the second injection as the gland is partially digested and may rupture with excessive pressure. After 10 min, the glands should be noticeably softer, indicating the success of the digestion. It is important to keep the collagenase incubation time to a minimum to avoid overdigestion of the medulla; 5. Open the gland longitudinally using a scalpel and clean scissors to reveal the cream-colored medulla; 6. Use scalpels to collect the medulla and transfer it to a clean Petri dish. Do not collect any purple material containing cortical cells. This procedure should be performed swiftly; 7. Mince the tissue into small pieces using two scalpels; 8. Transfer the material to the homemade filter (217-μm mesh); 9. Push the cells mechanically through the filter using a sterile glass rod of a Dounce homogenizer in Krebs balanced salt solution; 10. Collect the solution in 50-mL Falcon tubes and gently pellet the cells by centrifuging for 10 min at 100  g. Discard the supernatant and then resuspend the cell pellet in a final volume of 20 mL of Krebs balanced salt solution; 11. Filter cells by flow-gravity through a 70-μm mesh filter; 12. Collect about 19 mL of cell suspension, add 19 mL of the Percoll solution and gently mix; 13. Centrifuge at 20,000  g for 20 min at 24–26
C without braking to make the self-generated Percoll gradient [13, 14]. Be careful to equilibrate the tubes before spinning; 14. Discard 2–5 mL of the top gradient containing broken and cortical cells. Collect around 15–18 mL with a plastic Pasteur pipette at the gradient interface containing chromaffin cells and dispatch into four 50-mL Falcon tubes. Note that the bottom of the tubes contains red blood cells that should be discarded; 15. Complete each tube to 50 mL with DMEM-D and centrifuge for 10 min at 100  g at 20
C; 16. Resuspend the pellets in 10 mL of DMEM-D; 17. Count the cells in Trypan blue solution to estimate cell viability using a hemocytometer [15] (see Note 2). Prepare an aliquot of the cell sample for counting by adding 100 μL of 0.1% trypan blue stock solution (final concentration of 0.05% trypan blue)

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Fig. 1 Images of bovine chromaffin cells in culture. DIC images of bovine chromaffin cells after 4 (a) or 12 (b) days in culture. These cells display a round shape, with extensions after 12 days. Note that some cortical fibroblasts (flattened cells) can be found under the chromaffin cells after plating and will die progressively due to the use of antimitotic drugs in the medium. Bars ¼ 10 μm

and 80 μL of Krebs balanced salt solution to 20 μL of cell suspension in a 1.5-mL microcentrifuge tube. A typical yield is around 10–15  106 chromaffin cells per adrenal gland. 18. Plate the cells according to the requirements (see below) in DMEM-C and store for up to 2 weeks (Fig. 1). 3.2 Fluorescence Assay to Measure Catecholamine Secretion

Several companies sell kits to measure catecholamine levels developed for measurements of body fluid samples that are quite sensitive, but also rather tedious and expensive. The following assay is based on the oxidation of catecholamine into aminochrome [16], which is subsequently converted into aminolutine (Fig. 2). This method is much faster as it can be run in less than 2 h and is extremely cheap. The use of an adjustable volume multichannel pipette during the following steps minimizes pipetting and potential mistakes, but one should take care that each tip is always well adjusted to the holder. Each reagent is collected from a multichannel disposable solution basin that can be washed for reuse in a novel experiment. We recommend evaluating the sensitivity of the assay with chromaffin cells that you have prepared. 1. Plate freshly cultured chromaffin cells in 96-well plates at a density of 50,000; 100,000; or 200,000 cells in 200 μL of DMEM-D per well for 24 h to 48 h (see Note 3). In the following steps, the volumes of reagents or solutions are given for one well. 2. Place cells on a heated plate at 37
C and wash four times with 200 μL of Locke’s solution at 37
C for 10 min.

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Fig. 2 Principle of the fluorimetric catecholamine assay. Noradrenaline is synthesized from dopamine by dopamine β-hydroxylase (DBH) and further transformed into adrenaline by phenylethanolamine N-methyltransferase (PNMT). Catecholamine (adrenaline and noradrenaline) oxidation by sodium acetate, in the presence of potassium ferricyanide, leads to the production of aminochromes. Due to their unstable nature, these compounds are transformed into aminolutines in the presence of sodium hydroxide. These aminolutines emit at 520 nm when excited at 430 nm

3. Incubate with 50 μL of Locke’s solution (resting condition) or stimulate with Locke’s containing 10 μM of nicotine or highpotassium Locke’s solution at 37
C for 10 min. Typically, four to eight wells were used per condition, providing 12 to 24 different conditions in a 96-well plate. 4. Carefully collect the media containing the secreted material (50 μL) and briefly centrifuge for 5 min at 10,000  g to pellet cells that came off the plate. Store at 4
C. 5. Lyse the remaining cells by adding 200 μL of 0.1% Triton X-100 for 10 min at 37
C and pipetting up and down several times. Alternatively, three freeze and thaw cycles and careful shaking of the plate could also be used to lyse cells. 6. Dispatch 20 μL aliquots of secreted media and lysed cell solutions into two 96-well black-plates. 20 μL of Locke’s and 0.1% Triton X-100 solutions are also plated in one well of each plate to serve as blank reference.

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Fig. 3 Standard curves for adrenaline and noradrenaline detection using the fluorimetric catecholamine assay. Stock solutions of adrenaline, noradrenaline, and dopamine were serial-diluted and 20 μL aliquots were used for the fluorimetric catecholamine assay. The purple box indicates the concentration of catecholamine for which the assay is linear. n ¼ 8 samples per condition, and data are expressed as mean  S.E.M. (S.E.M. smaller than symbols are not shown)

7. Add successively 150 μL of 1 M sodium acetate and 15 μL of 0.25% potassium ferricyanide to each well to oxidize catecholamines to adrenochrome and noradrenochrome (Fig. 2). Shake gently for a couple of minutes. Formation of aminochromes occurs readily when catecholamines come into contact with Fe and display a deep red color. Because aminochromes are also quite unstable, they are not ideally suited for quantitative measurement. 8. Add 50 μL of 5 M sodium hydroxide to convert aminochromes to aminolutines (Fig. 2) [17]. 0.2% (w/v) ascorbic acid is included in the alkali reagent to prevent further oxidation (see Note 4). 9. Measure the fluorescence emitted by adrenolutine and noradrenolutine in secreted media (S) and in the cell lysates (P) (λex: 430 nm, λem: 520 nm) with a spectrofluorometer (see Note 4). Blank values (Locke’s and 0.1% Triton X-100 solutions) are subtracted from individual values obtained from the respective samples. Standard curves can be determined using known concentration of adrenaline and noradrenaline to demonstrate that the values obtained are in the linear range of detection of the

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assay (Fig. 3). In our hands, the assay is linear up to 200 ng/μL of catecholamines and is sensitive above 3 ng/μL (box in Fig. 3). It is of note that this assay does not display any significant sensitivity difference between adrenaline and noradrenaline, indicating that it is well suited to measure the secretory activity of mixed adrenergic and noradrenergic cell population. Moreover, this assay does not detect dopamine even at high levels and as such is not adapted for PC12 cells, which do not express dopamine β-hydroxylase. Note that in order to avoid variations originating from differences in cell quantities per well, the amount of catecholamine released (X) is better expressed as a percentage of the total amount of catecholamine present in cells before stimulation using the following formula: Catecholamine release ð%of totalÞ ¼

S  2:5  100 ðS  2:5Þ þ ðP  10Þ

S: value of one individual secreted medium aliquot from which the blank value of Locke’s solution has been subtracted. P: value of the corresponding cell aliquot from which the blank value of the 1% Triton X-100 solution has been subtracted. Among the secretagogues tested here, nicotine is more efficient than high potassium to trigger catecholamine secretion from bovine chromaffin cells (Fig. 4). The secretion levels observed using this assay are very similar to those obtained with [3H]noradrenaline [18–21]. We also found that the levels of basal release measured from 50,000 cells are usually close to the detection limit and therefore less accurately estimated. In conclusion, using 100,000 or 200,000 cells per well appears well adapted for this secretion assay.

4

Notes 1. The isolation procedure should focus first on obtaining healthy cells rather than the highest yield. We recommend adjusting the concentration of collagenase used for digestion and minimizing the mechanical disruption during tissue digestion. 2. A standard Neubauer hemocytometer remains a valuable and cheap tool to count cells and to get a general idea of their viability and contamination with erythrocytes. We prepare a 1:9 dilution of cells by mixing 20 μL of cell suspension plus 100 μL of Krebs balanced salt solution plus 80 μL of trypan blue staining solution (see above). Guidelines for the correct use of hemocytometers have been published previously [15].

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Fig. 4 Secretion of catecholamines triggered by nicotine or high potassium and measured by fluorimetric catecholamine assay. In a 96-well plate, 50,000, 100,000, or 200,000 cells (numbers depicted in x axis) were seeded per well for 24 h. After four washes with Locke’s solution, cells were incubated for 10 min at 37
C in Locke’s solution (resting) or stimulated with Locke’s solution containing 10 μM of nicotine or 59 mM potassium solution. Catecholamine levels secreted in the medium and remaining in cells were estimated using a fluorimetric assay. The secretory activity is expressed as a percentage of the total amount of catecholamine present in cells. Data are expressed as the mean  S.E.M. (n ¼ 3)

3. Although cells can be maintained for over a week after plating, optimal secretion usually occurs within 24–48 h of plating. However, we noticed that in some rare cases, nicotine was unable to trigger catecholamine release, especially in the first days of culture, most likely because of partial degradation of the nicotinic receptor by collagenase. 4. Note that the fluorescence will slowly decay with time, so it is best to take the reading as soon as the assay is finished.

Acknowledgments We thank Dr. F.A. Meunier (University of Queensland, Brisbane, Australia) and Dr. Robert D. Burgoyne (University of Liverpool) for sharing an original fluorescence assay for catecholamine secretion. We are indebted to the Abattoir Municipal of Haguenau (France) for providing us with the fresh bovine adrenal glands. This work was supported by Fondation pour la Recherche Me´dicale and the ANR grant (ANR-19-CE44-0019) to NV.

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References 1. Bader MF, Holtz RW, Kumakura K, Vitale N (2002) Exocytosis: the chromaffin cell as a model system. Ann N Y Acad Sci 971:178–183 2. Vitale N, Gensse M, Chasserot-Golaz S, Aunis D, Bader MF (1996) Trimeric G proteins control regulated exocytosis in bovine chromaffin cells: sequential involvement of go associated with secretory granules and Gi3 bound to the plasma membrane. Eur J Neurosci 8:1275–1285 3. Zeniou-Meyer M, Liu Y, Be´gle´ A, Olanich ME, Hanauer A, Becherer U, Rettig J, Bader MF, Vitale N (2008) The coffin-Lowry syndromeassociated protein RSK2 is implicated in calcium-regulated exocytosis through the regulation of PLD1. Proc Natl Acad Sci U S A 105:8434–8439 4. Be´gle´ A, Tryoen-To´th P, de Barry J, Bader MF, Vitale N (2009) ARF6 regulates the synthesis of fusogenic lipids for calcium-regulated exocytosis in neuroendocrine cells. J Biol Chem 284:4836–4845 5. Domı´nguez N, Rodrı´guez M, Machado JD, Borges R (2012) Preparation and culture of adrenal chromaffin cells. Methods Mol Biol 846:223–234 6. Kloppenborg PW, Island DP, Liddle GW, Michelakis AM, Nicholson WE (1968) A method of preparing adrenal cell suspensions and its applicability to the in vitro study of adrenal metabolism. Endocrinology 82:1053–1058 7. Hochman J, Perlman RL (1976) Catecholamine secretion by isolated adrenal cells. Biochim Biophys Acta 421:168–175 8. O’Connor DT, Mahata SK, Mahata M, Jiang Q, Hook VY, Taupenot L (2007) Primary culture of bovine chromaffi n cells. Nat Protoc 2:1248–1253 9. Krause W, Michael N, Lubke C, Livett BG, Oehme P (1996) Catecholamine release from fractionated chromaffin cells. Eur J Pharmacol 302:223–228 10. Livett BG, Boksa P, Dean DM, Mizobe F, Lindenbaum MH (1983) Use of isolated chromaffin cells to study basic release mechanisms. J Auton Nerv Syst 7:59–86 11. Bader MF, Ciesielski-Treska J, Thierse D, Hesketh JE, Aunis D (1981) Immunocytochemical study of microtubules in chromaffi n cells in culture and evidence that tubulin is not an

integral protein of the chromaffin granule membrane. J Neurochem 37:917–933 12. Baker PF, Knight DE (1981) Calcium control of exocytosis and endocytosis in bovine adrenal medullary cells. Philos Trans R Soc Lond Ser B Biol Sci 296:83–103 13. Bader MF, Trifaro´ JM, Langley OK, Thierse´ D, Aunis D (1986) Secretory cell actin-binding proteins: identification of a gelsolin-like protein in chromaffin cells. J Cell Biol 102:636–646 14. Bader MF, Thierse´ D, Aunis D, Ahnert-HilgerG, Gratzl M (1986) Characterization of hormone and protein release from alpha-toxinpermeabilized chromaffin cells in primary culture. J Biol Chem 261:5777–5783 15. Strober W (2001) Trypan blue exclusion test of cell viability. Curr Protoc Immunol. Appendix 3, Appendix 3B 16. Meunier FA, Feng ZP, Molgo J, Zamponi GW, Schiavo G (2002) Glycerotoxin from Glycera convoluta stimulates neurosecretion by up-regulating N-type Ca2+ channel activity. EMBO J 21:6733–6743 17. Heacock RA, Laidlaw BD (1958) Reduction of adrenochrome with ascorbic acid. Nature 182:526–527 18. Vitale N, Mukai H, Rouot B, Thierse´ D, Aunis D, Bader MF (1993) Exocytotis in chromaffin cells. Possible involvement of the heterotrimeric GTP-binding protein Go. J Biol Chem 268:14715–14723 19. Chasserot-Golaz S, Vitale N, Umbrecht-JenckE, Knight D, Gerke V, Bader MF (2005) Annexin 2 promotes the formation of lipid microdomains required for calcium-regulated exocytosis of dense-core vesicles. Mol Biol Cell 16:1108–1119 20. Tryoen-To´th P, Chasserot-Golaz S, Tu A, Gherib P, Bader MF, Beaumelle B, Vitale N (2013) HIV-1 Tat protein inhibits neurosecretion by binding to phosphatidylinositol 4,5-bisphosphate. J Cell Sci 126:454–463 21. Gabel M, Delavoie F, Royer C, Thahouly T, Gasman S, Bader MF, Vitale N, ChasserotGolaz S (2019) Phosphorylation cycling of Annexin A2 Tyr23 is critical for calciumregulated exocytosis in neuroendocrine cells. Biochim Biophys Acta, Mol Cell Res 1866:1207–1217

Chapter 12 Measurement of Exocytosis in Genetically Manipulated Mast Cells Ofir Klein, Nurit P. Azouz, and Ronit Sagi-Eisenberg Abstract The hallmark of mast cell activation is secretion of immune mediators by regulated exocytosis. Measurements of mediator secretion from mast cells that are genetically manipulated by transient transfections provide a powerful tool for deciphering the underlying mechanisms of mast cell exocytosis. However, common methods to study regulated exocytosis in bulk culture of mast cells suffer from the drawback of high signal-to-noise ratio because of their failure to distinguish between the different mast cell populations, that is, genetically modified mast cells versus their non-transfected counterparts. In particular, the low transfection efficiency of mast cells poses a significant limitation on the use of conventional methodologies. To overcome this hurdle, we developed a method, which discriminates and allows detection of regulated exocytosis of transfected cells based on the secretion of a fluorescent secretory reporter. We used a plasmid encoding for Neuropeptide Y (NPY) fused to a monomeric red fluorescent protein (NPY-mRFP), yielding a fluorescent secretory granule-targeted reporter that is co-transfected with a plasmid encoding a gene of interest. Upon cell trigger, NPY-mRFP is released from the cells by regulated exocytosis, alongside the endogenous mediators. Therefore, using NPY-mRFP as a reporter for mast cell exocytosis allows either quantitative, via a fluorimeter assay, or qualitative analysis, via confocal microscopy, of the genetically manipulated mast cells. Moreover, this method may be easily modified to accommodate studies of regulated exocytosis in any other type of cell. Key words Mast cells, Regulated exocytosis, NPY-mRFP, Transfection, Microscopy

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Introduction Mast cells (MCs) are granulocyte cells of the immune system that originate in the bone marrow and reach full maturation upon homing to connective and mucosal tissues [1]. The hallmark of MC function in either their innate or adaptive immune responses is their fast release of inflammatory mediators and proteases, which are prestored in secretory granules (SGs), by utilizing different modes of regulated exocytosis [2]. MCs release the contents of their SGs either directly after encountering an allergen, in a process that involves the formation of allergen-specific immunoglobulin E

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(IgE) antibodies, or indirectly of IgE, by numerous stimuli, including neuropeptides and toxins [3]. Following their exocytosis, MC inflammatory mediators affect multiple targets and thereby induce allergic symptoms such as pain, high fever, difficulty in breathing, and hypotension. The severity of allergic reactions may range from local discomfort in cases such as skin rash to death by anaphylaxis, defined by the World Health Organization as a severe, lifethreatening, generalized or systemic hypersensitivity reaction [4]. Therefore, measurement of regulated exocytosis in MCs is not only important for delineating the underlying mechanisms of MC secretion in basic and clinical research of MC physiology in health and disease, but also as a potential therapeutic target to control allergic diseases. Many different secretory reporters have been established in MCs in order to quantify regulated exocytosis. The latter include endogenous mediators, such as histamine, and proteases, such as β-hexosaminidase, whose release from activated cells is most commonly monitored, as a measure of exocytosis [5–14], and exogenous reporters, such as fluorescently labeled dextran or avidin, that are added to the media and taken up by the cells [15–22], or overexpression of pHluorin-tagged β-hexosaminidase [23]. The detection of these reporters is achieved by colorimetric or fluorometric assays, radiolabeling methods, radioimmunoassays, and microscopy-based methods in vitro and in vivo [5–23]. MC secretion assays are often coupled with genetic modifications, which modulate the cell’s responsiveness, in order to explore the impact of overexpression, mutation, or knockdown of target genes on exocytosis. However, MCs are difficult to transfect, and hence, it is challenging to perform a large-scale screen for the effect of genetic manipulations on MC secretion, when only a small fraction of cells is transfected. For this purpose, we devised a method based on the co-transfection of plasmids encoding genes of interest, or their targeting shRNAs, with a plasmid encoding a secretory reporter gene, that is, Neuropeptide Y fused to a monomeric red fluorescent protein (NPY-mRFP) [24]. We have shown that this reporter fulfills the properties that are required of a genuine reporter for secretion, namely it localizes to the SGs of the MCs and is secreted from the cells alongside the endogenous inflammatory mediators [24]. Furthermore, unlike pHluorin, which was previously used as a transfectable fluorescent MC SG reporter, when fused to β-hexosaminidase [23], and which is pH-sensitive and therefore quenched when packed inside the acidic SGs of the MCs [25], mRFP is pH-insensitive. Therefore, while the use of pHluorin as a fluorescent marker allows the specific detection of live events of exocytosis [23, 26–29], an mRFP-based reporter can additionally serve as a suitable probe for detecting SG dynamics inside the cell.

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Fig. 1 Scheme of transient transfection and NPY-mRFP detection. (a) Cells are mixed with a large amount of a modulating (tested) gene plasmid (blue) and a small amount of NPY-mRFP reporter plasmid (red) and electroporated. (b) As a result, while some of the cells took up only the modulating plasmid, all the cells, which took up the NPY-mRFP reporter plasmid, have also taken up the modulating plasmid. (c) Analysis of NPY-mRFP secretion by either a fluorescence plate reader (quantitative) or a fluorescence microscope (qualitative) reflects the secretion exclusively from the genetically manipulated cells that co-express the modulating gene, regardless of the transfection efficiency

Hence, the method described here relies on: (1) the pH-insensitive secretory reporter NPY-mRFP, which enables the imaging of non-secreting SG dynamics, and (2) the co-transfection of cells to co-express the reporter gene with the modulating gene (i.e., overexpression of a wild-type or a mutant gene, or a gene targeting shRNA), allowing for quantitative and selective measurement of exocytosis of the transfected cells within a heterogenous population of cells. To ensure the high efficiency of NPY-mRFP incorporation into cells that are expressing the modulating gene, the co-transfection consists of the application of a higher amount of the plasmid that encodes the modulating gene than the amount of the NPY-mRFP reporter plasmid, boosting the probability of co-expression of NPY-mRFP and the modulating gene. Furthermore, visualization of the cells that carry the mRFP fluorescent signal under the microscope reveals the phenotype of the cells that also express the modulating gene, thus allowing not only functional screening of exocytosis, but also phenotypic screening of MCs (Fig. 1). While the method presented here is calibrated to study regulated exocytosis in MCs by the detection of NPY in cell supernatants, it may be applied to many other cell types. NPY is a

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highly conserved, low-molecular weight secreted peptide, which has been found to localize in SGs in different cell types [26, 27, 30], including cells of the nervous system [28, 31]. Moreover, this method may also be customized for any type of cell, simply by tagging a protein, distinct from NPY, that is secreted in a regulated fashion in the desired cell type. Moreover, the method may be modified to study other forms of exocytosis, such as continuous or nonconventional exocytosis, by tagging an adequate cargo protein with a fluorescent marker (see Note 1).

2

Materials 1. Culture and Transfection of RBL-2H3 (A Prevalent MC Model, Hereafter Referred to as RBL) Cells. (a) RBL cells (see Note 2). (b) RBL cell growth media: low-glucose (1000 mg/L) Dulbecco’s modified Eagle medium (DMEM) supplemented with 10% FBS, 100 μg/mL streptomycin, 100 units/mL penicillin, 12 units/mL nystatin and 2 mM L-glutamine. (c) Trypsin/EDTA solution B. (d) Standard tissue-treated maintenance).

10-cm

dishes

(for

cell

(e) Chambered coverglass or glass bottom culture dishes (for live cell microscopy). (f) Standard 24-well, flat-bottom cell culture-treated plates (for quantitative secretion assay). (g) 4-mm electroporation cuvettes. (h) Transfection buffer: 20 mM K-Pipes pH 7, 10 μM Ca2+ acetate, 2 mM Mg2+ acetate, 128 mM potassium glutamate in low-glucose DMEM (1000 mg/L) (see Note 3). (i) Purified NPY-mRFP (reporting) plasmid and modulating plasmid at high concentration (at least 1 μg/μL). (j) Square current electroporator. 2. Mast cell activation (for either quantitative or qualitative assays). (a) Tyrode’s buffer: 2.7 mM KCl, 1 mM MgCl2, 137 mM NaCl, 0.4 mM NaH2PO4, 1.8 mM CaCl2, 0.1% BSA, 5.6 mM glucose and 20 mM HEPES (pH 7) in doubledistilled water (DDW) (see Note 4). (b) Allergen: DNP-HSA—prepare a 20
working solution (1 μg/mL) from a stock solution of 10 mg/mL and keep on ice, shielded from light. (c) IgE anti-DNP, clone SPE-7.

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3. Quantitative Measurement of NPY-mRFP Release. (a) 96-well, flat-bottom plates. (b) Fluorescence plate reader. (c) Lysis buffer—Tyrode’s buffer supplemented with 0.5% Triton X-100. 4. Imaging of NPY-mRFP Release. (a) Confocal fluorescence microscope equipped with a CO2 controller (4.8%), heated chamber (37  C) and a C-Apochromat
63/1.2 W Corr objective (see Note 5).

3

Methods MCs can be triggered for secretion by different stimuli. Here we describe MC activation by the classic allergic pathway, which requires sensitization of the Fc-epsilon RI receptor by incubation of the cells with allergen-specific IgE (herein DNP-specific IgE) prior to activation with the allergen (DNP-HSA, hereafter referred to as Ag). Other popular mechanisms for triggering MCs for secretion include activation of the MRGPRX2 receptor by different ligands [32] or synthetic activation by the combination of Phorbol 12-myristate 13-acetate and calcium ionophore [9]. MCs of primary or cell line origin vary in their adherence properties when grown in culture, but they are all-incompatible with transfection by either calcium or lipofectamine. Here we describe transfection of RBL cells, which are adherent and most commonly transfected by electroporation. However, this method may be calibrated to fit any other cell type, by adjusting the growth and transfection conditions.

3.1 RBL Cell Line Transfection

3.1.1 Transfection of RBL Cells

One transfection reaction of RBL cells requires 1.5
107 cells. Therefore, 1 day prior to the transfection day, seed 2.5
106 RBL cells in two 10-cm culture plates per transfection, to reach a maximum confluency of 70–80% on the day of transfection. 1. To detach RBL cells from the culture dishes, aspirate the media and replace with 2 mL of trypsin/EDTA per culture dish. 2. Incubate at 37  C until cells detach (but not over 10 min). 3. Add 2 mL of culture medium to each plate in order to neutralize the trypsin solution and transfer the cells from two 10-cm culture plates to a single 15-mL conical tube. 4. Centrifuge for 5 min at 300
g, 25  C to sediment the cells. 5. Aspirate the supernatant carefully, so as to not dislodge the cell pellet.

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Fig. 2 Plate setup example. (a) Duplicates of cells transfected with NPY-mRFP and a control empty vector are seeded in columns 1–2, and duplicates of cells transfected with NPY-mRFP and the modulating gene plasmid are seeded in columns 3–4. Non-transfected, untreated cells are seeded in columns 5–6. Row A is to be used as an untreated control, while rows B–D may be used for different activation treatments such as time/dose responses or treatment with a drug. (b) After cell activation in the 24-well plate, the cell supernatants, that contain the secreted NPY-mRFP, are transferred to the corresponding columns 1–4 in a 96-well plate (EV ¼ empty vector, MG ¼ modulating gene). Then, the cell lysates (made in Tyrode’s buffer supplemented with 0.5% Triton X-100), containing the retained, non-secreted NPY-mRFP are transferred to the corresponding columns 5–8 of the 96-well plate. Supernatants derived from non-transfected cells serve as blanks and are placed in wells A9 and A10 (NT sup). Their lysates (made in Tyrode’s buffer supplemented with 0.5% Triton X-100) are placed in duplicate in wells A11 and A12, serving as blanks for the cell lysates (NT lysate)

6. Resuspend the cell pellet in 380 μL transfection buffer and transfer to a 4-mm cuvette. 7. Add up to 20 μg of NPY-mRFP and up to 30 μg of plasmid of choice or control empty vector, resulting in a maximum of 50 μg plasmids (see Notes 6 and 7). It is recommended to use a minimal volume of plasmids in order to maintain the transfection buffer properties (see Note 8). 8. Place the cuvette on ice for 10 min. 9. With a paper towel, wipe off any excess moist from the cuvette, place it in the electroporator and pulse at 300 V for 20 ms. 10. Immediately, seed the cells into prewarmed culture media containing 250 ng/mL of IgE: (a) For quantitative measurement of exocytosis, seed 8 μL (~4
105 cells) of electroporated cells in 24-well plates containing 500 μL of IgE-supplemented media and incubate overnight. Prepare wells in duplicates or triplicates for each treatment, including untreated controls. For blank controls, seed 4
105 cells of non-transfected cells (Fig. 2).

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(b) For live cell imaging, seed 1 μL (~0.5
105 cells) of electroporated cells in eight-well chambers containing 300 μL of IgE-supplemented media and incubate overnight. 1. After 24–48 h of transfection, aspirate media from the wells and wash three times with Tyrode’s buffer.

3.2 Quantitative Measurement of NPY-mRFP Release

2. Immerse non-transfected control (blanks) and untreated control wells in 200 μL of Tyrode’s buffer and experimental wells in 190 μL of Tyrode’s buffer.

3.2.1 Activation of RBL Cells

3. To activate cells for exocytosis, add 10 μL of 20
Ag solution to each of the experimental wells and incubate for the desired time periods at 37  C. 1. To collect the secreted NPY-mRFP, transfer 180 μL from each well to a flat-bottom 96-well plate, suitable for a fluorescence plate reader. It is important to transfer the supernatant gently in order to avoid bubbles, which can interfere with fluorescence reading (see Note 9) (see Fig. 2 for plate setup example).

3.2.2 Sample Collection of Secreted and Non-secreted Fractions

2. Aspirate the remaining media from the 24-well plate and replenish with 200 μL of Tyrode’s buffer supplemented with 0.5% Triton X-100 and incubate at 37  C for 10 min to lyse the cells. 3. To collect the non-secreted NPY-mRFP reservoir of the cell, repeat step 1 (see Fig. 2 for plate setup example). 4. Measure mRFP fluorescence using a fluorescence plate reader using the appropriate filter setting available. mRFP excitation peaks at 584 nm and emission peaks at 607 nm [33]. It is recommended not to exceed excitation/emission parameters by more than 20 nm of the peaks. 5. To calculate percentage of secretion, first average duplicates. Then, subtract the supernatant blank value from the readings of the supernatant wells, and the lysate blank value from the lysate well readings. Then, calculate percentage of NPY-mRFP secretion as follows (for a detailed example, see Fig. 3): % secreted NPY‐mRFP ¼

3.3 Live Cell Imaging of NPY-mRFP Release

supernatant value
100 ðsupernatant value þ lysate valueÞ

1. After 24–48 h of transfection, wash the cells three times with prewarmed Tyrode’s buffer and replenish each chamber with 285 μL of Tyrode’s buffer. 2. Place the eight-well chambered coverglass and secure it in place in the microscope’s CO2/temperature-controlled incubator. 3. Select the appropriate excitation/emission settings available on the microscope, which suit the fluorophores used in the

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Fig. 3 Example of secretion calculation. Data from a typical experiment of RBL cells that were co-transfected with 15 μg of a plasmid encoding NPY-mRFP and 30 μg of a plasmid encoding EGFP. The cells were sensitized with IgE and activated on the following day with Ag for increasing time periods. NPY-mRFP levels were assayed and the extent of secretion quantified as detailed in methods. (a) Average NPY-mRFP fluorescence values were calculated from duplicate fluorescence values of the supernatants and lysates of the tested wells and the non-transfected blank controls (NT). (b) In order to correct the fluorescence values for the autofluorescence of the buffers and the cells, the average NT sup value (i.e., 52.7) was deducted from the average supernatant values of the tested wells and the average NT lysate value (i.e., 56) was deducted from the average lysate values of the tested wells. (c) To calculate the percentage of NPY-mRFP secretion at each time point, the corrected supernatant values shown in (b) were divided by the sum of the corrected supernatant and lysate values from (b) and multiplied by 100. (d) A scatter line graph representing the percentage of secretion as function of time

experiment. For mRFP, excitation peak is at 584 nm and emission peak is at 607 nm (see Note 10). 4. Select minimal excitation level and minimal exposure time to reduce bleaching and cell toxicity during live cell imaging and the appropriate interval time between acquisitions (see Note 11). 5. Locate the cells for imaging and set the desired zoom and focus. 6. Begin image acquisition prior to cell activation to obtain the steady-state dynamics of the SGs in the cell. 7. To activate the cells, add 15 μL of Ag solution. Be cautious not to move the chamber so that the cells’ position does not change. If needed, quickly adjust the chamber’s position and focus after addition of Ag. 8. Continue imaging for 15 min (see Note 11).

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Notes 1. In MCs, as in many other secretory cells, the SGs are acidic. Therefore, when choosing a fluorescent marker, it is recommended to use a pH-insensitive fluorescent protein, such as mRFP, which retains its fluorescence in an acidic environment. 2. We chose to use RBL cells, a reliable MC model, since they are relatively easy to handle and image [34]. 3. Transfection buffer may be prepared in advance and aliquoted and stored at 20  C. 4. A 20
Tyrode salt solution (54 mM KCl, 20 mM MgCl2, 2.74 M NaCl, and 8 mM NaH2PO4 in DDW) may be prepared in advance and stored at 4  C. To make the final Tyrode’s buffer, dilute the solution in DDW and supplement with 0.1% BSA, 1.8 mM CaCl2, 5.6 mM (0.1%) glucose, and 20 mM HEPES (pH 7). 5. Instead of a standard confocal microscope, total internal reflection microscope (TIRFM) may also be used, for a more sensitive detection of the fusion event of the SG with the plasma membrane [35, 36]. 6. When using a modulating plasmid that is larger than the reporter plasmids, it is important to remember to maintain a molar ratio in favor of the modulating plasmid (at least 1:1.5). When using significantly larger plasmids (10 kilobase and more), whose transfection efficiency is likely to be lower, it is also recommended to increase the molar ratio further, in favor of the modulating plasmid. 7. We routinely use up to three plasmids in one transfection, allowing for a combination of two modifying plasmids and the SG reporter. Under such circumstances, we recommend using a ratio of 10 μg of reporter plasmid, 15 μg of modulating plasmid #1 and 20 μg of modulating plasmid #2, in order to ensure that the cells which took up the reporter plasmid also took up both modulating plasmids. Therefore, under such conditions, the transfection will yield cells transfected with modulating plasmid #2, cells transfected with modulating plasmids #1 and #2, and cells transfected with modulating plasmid #1, #2, and the reporter plasmid, ensuring that indeed the reporter plasmid reports only for cells that co-express the two modulating plasmids. 8. For efficient transfection, it is imperative to maintain the osmolarity and the conductivity of the transfection buffer [37, 38]. Therefore, it is imperative to use a highconcentration plasmid prep in order not to overdilute the transfection media when adding the plasmids.

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9. In case there are bubbles, burst them by pricking with a needle. 10. Most modern fluorescence microscopes are equipped with a built-in setting for common fluorophores. It is usually possible to choose the recommended settings for optimal detection of the fluorophores to be used from a list of fluorophores that is included in the microscope’s software. 11. It is important to minimize exposure of the cells to the laser in order to reduce cell toxicity and avoid bleaching of the fluorophores. Therefore, both the laser power and exposure time should be set to the possible minimum. The reduction of the exposure time can be achieved by increasing the scan speed and binning. Another important parameter affecting the total laser exposure time is the time interval between the image acquisitions. For fast secreting cells, such as the MCs, the interval time should be limited to a few seconds. However, since the kinetics of secretion differs between different cell types [39], the acquisition interval and overall duration of acquisition should be calibrated and adjusted to each cell type. References 1. Dahlin JS, Hallgren J (2015) Mast cell progenitors: origin, development and migration to tissues. Mol Immunol 63:9–17. https:// doi.org/10.1016/j.molimm.2014.01.018 2. Klein O, Sagi-Eisenberg R (2019) Anaphylactic degranulation of mast cells: focus on compound exocytosis. J Immunol Res 2019:1–12. https://doi.org/10.1155/2019/9542656 3. Wernersson S, Pejler G (2014) Mast cell secretory granules: armed for battle. Nat Rev Immunol 14:478–494. https://doi.org/10. 1038/nri3690 4. Johansson SGO, Bieber T, Dahl R et al (2004) Revised nomenclature for allergy for global use: report of the nomenclature review Committee of the World Allergy Organization, October 2003. J Allergy Clin Immunol 113:832–836. https://doi.org/10.1016/j. jaci.2003.12.591 5. Ra˚dinger M, Jensen BM, Swindle E, Gilfillan AM (2015) Assay of mast cell mediators. In: Methods in molecular biology. Humana Press, New Jersey, pp 307–323 6. Cruse G, Gilfillan AM, Smrz D (2015) Flow cytometry-based monitoring of mast cell activation. In: Methods in molecular biology. Humana Press, New Jersey, pp 365–379 7. Johnson DA (2006) Human mast cell proteases: activity assays using thiobenzyl ester substrates. In: Mast cells. Humana Press, New Jersey, pp 193–202

8. Chi DS, Fitzgerald SM, Krishnaswamy G (2006) Mast cell histamine and cytokine assays. In: Mast cells. Humana Press, New Jersey, pp 203–216 9. Jensen BM, Falkencrone S, Skov PS (2014) Measuring histamine and cytokine release from basophils and mast cells. In: Methods in molecular biology. Humana Press, New Jersey, pp 135–145 10. Hohman RJ, Dreskin SC (2001) Measuring degranulation of mast cells. In: Current protocols in immunology. John Wiley & Sons, Inc., Hoboken, NJ, USA, p Unit 7.26 11. Kuehn HS, Radinger M, Gilfillan AM (2010) Measuring mast cell mediator release. In: Current protocols in immunology. John Wiley & Sons, Inc., Hoboken, NJ, USA, p Unit7.38 12. Demo SD, Masuda E, Rossi AB et al (1999) Quantitative measurement of mast cell degranulation using a novel flow cytometric annexinV binding assay. Cytometry 36:340–348. https://doi.org/10.1002/(SICI)1097-0320( 19990801)36:43.0. CO;2-C 13. Naal RMZG, Tabb J, Holowka D, Baird B (2004) In situ measurement of degranulation as a biosensor based on RBL-2H3 mast cells. Biosens Bioelectron 20:791–796. https://doi. org/10.1016/J.BIOS.2004.03.017 14. Gadi D, Wagenknecht-Wiesner A, Holowka D, Baird B (2011) Sequestration of

Exocytosis in Transiently Transfected Mast Cells phosphoinositides by mutated MARCKS effector domain inhibits stimulated Ca2+ mobilization and degranulation in mast cells. Mol Biol Cell 22:4908–4917. https://doi.org/10. 1091/mbc.e11-07-0614 15. Cohen R, Holowka DA, Baird BA (2015) Realtime imaging of Ca(2+) mobilization and degranulation in mast cells. In: Methods in molecular biology. Humana Press, New Jersey, pp 347–363. https://doi.org/10.1007/9781-4939-1568-2_22 16. Gaudenzio N, Sibilano R, Marichal T et al (2016) Different activation signals induce distinct mast cell degranulation strategies. J Clin Invest 126:3981–3998. https://doi.org/10. 1172/JCI85538 17. Klein O, Roded A, Hirschberg K et al (2018) Imaging FITC-dextran as a reporter for regulated exocytosis. J Vis Exp 136:57936. https:// doi.org/10.3791/57936 18. Williams RM, Webb WW (2000) Single granule pH cycling in antigen-induced mast cell secretion. J Cell Sci 113(Pt 21):3839–3850 19. Williams RM, Shear JB, Zipfel WR et al (1999) Mucosal mast cell secretion processes imaged using three-photon microscopy of 5-hydroxytryptamine autofluorescence. Biophys J 76:1835–1846. https://doi.org/10. 1016/S0006-3495(99)77343-1 20. Kawasaki Y, Saitoh T, Okabe T et al (1991) Visualization of exocytotic secretory processes of mast cells by fluorescence techniques. Biochim Biophys Acta Biomembr 1067:71–80. https://doi.org/10.1016/0005-2736(91) 90027-6 21. Balseiro-Gomez S, Flores JA, Acosta J et al (2016) Transient fusion ensures granule replenishment to maintain repeated release after IgE-mediated mast cell degranulation. J Cell Sci 129:3989–4000. https://doi.org/10. 1242/jcs.194340 22. Tharp MD, Seelig LL, Tigelaar RE, Bergstresser PR (1985) Conjugated avidin binds to mast cell granules. J Histochem Cytochem 33:27–32. https://doi.org/10.1177/33.1. 2578142 23. Wilson JD, Shelby SA, Holowka D, Baird B (2016) Rab11 regulates the mast cell exocytic response. Traffic 17:1027–1041. https://doi. org/10.1111/tra.12418 24. Azouz NP, Matsui T, Fukuda M, SagiEisenberg R (2012) Decoding the regulation of mast cell exocytosis by networks of Rab GTPases. J Immunol 189:2169–2180. https://doi.org/10.4049/jimmunol. 1200542

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25. Blott EJ, Griffiths GM (2002) Secretory lysosomes. Nat Rev Mol Cell Biol 3:122–131. https://doi.org/10.1038/nrm732 26. Makhmutova M, Liang T, Gaisano H et al (2017) Confocal imaging of neuropeptide Y-pHluorin: a technique to visualize insulin granule exocytosis in intact murine and human islets. J Vis Exp:e56089. https://doi. org/10.3791/56089 27. Almac¸a J, Liang T, Gaisano HY et al (2015) Spatial and temporal coordination of insulin granule exocytosis in intact human pancreatic islets. Diabetologia 58:2810–2818. https:// doi.org/10.1007/s00125-015-3747-9 28. Dominguez N, van Weering JRT, Borges R et al (2017) Dense-core vesicle biogenesis and exocytosis in neurons lacking chromogranins A and B. J Neurochem 144:241–254. https:// doi.org/10.1111/jnc.14263 29. Burrone J, Li Z, Murthy VN (2007) Studying vesicle cycling in presynaptic terminals using the genetically encoded probe synaptopHluorin. Nat Protoc 1:2970–2978. https://doi. org/10.1038/nprot.2006.449 30. Jacobs DT, Weigert R, Grode KD et al (2009) Myosin Vc is a molecular motor that functions in secretory granule trafficking. Mol Biol Cell 20:4471–4488. https://doi.org/10.1091/ mbc.e08-08-0865 31. Tan CMJ, Green P, Tapoulal N et al (2018) The role of neuropeptide Y in cardiovascular health and disease. Front Physiol 9:1281. https://doi.org/10.3389/fphys.2018.01281 32. Redegeld FA, Yu Y, Kumari S et al (2018) Non-IgE mediated mast cell activation. Immunol Rev 282:87–113. https://doi.org/10. 1111/imr.12629 33. Campbell RE, Tour O, Palmer AE et al (2002) A monomeric red fluorescent protein. Proc Natl Acad Sci 99:7877–7882. https://doi. org/10.1073/pnas.082243699 34. Falcone FH, Wan D, Barwary N, SagiEisenberg R (2018) RBL cells as models for in vitro studies of mast cells and basophils. Immunol Rev 282:47–57. https://doi.org/ 10.1111/imr.12628 35. Klein O, Krier-Burris RA, Lazki-Hagenbach P et al (2019) Mammalian diaphanous-related formin 1 (mDia1) coordinates mast cell migration and secretion through its actin-nucleating activity. J Allergy Clin Immunol 144 (4):1074–1090. https://doi.org/10.1016/j. jaci.2019.06.028 36. Ljubicic S, Bezzi P, Brajkovic S et al (2013) The GTPase rab37 participates in the control of insulin exocytosis. PLoS One 8:e68255.

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38. Luft C, Ketteler R (2015) Electroporation knows no boundaries: the use of electrostimulation for siRNA delivery in cells and tissues. J Biomol Screen 20:932–942. https://doi.org/ 10.1177/1087057115579638 39. Martin TFJ (2003) Tuning exocytosis for speed: fast and slow modes. Biochim Biophys Acta, Mol Cell Res 1641:157–165. https:// doi.org/10.1016/S0167-4889(03)00093-4

Chapter 13 Super-Resolution Microscopy and Particle-Tracking Approaches for the Study of Vesicular Trafficking in Primary Neutrophils Jennifer L. Johnson, Kersi Pestonjamasp, William B. Kiosses, and Sergio D. Catz Abstract Neutrophils are short-lived cells after isolation. The analysis of neutrophil vesicular trafficking requires rapid and gentle handling. Recently developed super-resolution microscopy technologies have generated unparalleled opportunities to help understand the molecular mechanisms regulating neutrophil vesicular trafficking, exocytosis, and associated functions at the molecular level. Here, we describe super-resolution and total internal reflection fluorescence (TIRF) microscopy approaches for the analysis of vesicular trafficking and associated functions of primary neutrophils. Key words Total internal reflection fluorescence (TIRF), Vesicular trafficking, Neutrophils

1

Introduction The study of neutrophil cell biology is largely limited by their short life after isolation and requires gentle handling when manipulating these cells to avoid activation. Neutrophils have several secretory organelles that regulate the timely response of these cells to infections and proinflammatory cues [1]. The analysis of the distribution of trafficking molecules that control these organelles has led to new understandings of how these mechanisms regulate neutrophil function. A limitation of super-resolution techniques is that the interpretation of the molecular distribution of individual trafficking markers is restricted by the density at which these molecules populate a given organelle. Thus, while reconstruction and localization are relatively easy for molecules that are present in such organelles in high density, the task becomes more difficult for molecules whose distribution at selective organelles is sparse. Here, we describe analytical and quantitative methods that overcome this

Florence Niedergang, Nicolas Vitale and Ste´phane Gasman (eds.), Exocytosis and Endocytosis: Methods and Protocols, Methods in Molecular Biology, vol. 2233, https://doi.org/10.1007/978-1-0716-1044-2_13, © Springer Science+Business Media, LLC, part of Springer Nature 2021

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limitation and establish methods to calculate the relative proximity of molecular interactors at neutrophil secretory granules.

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Materials and Equipment 1. Phenol Red-Free Roswell Park Memorial Institute (RPMI) 1640, Medium (PRF-RPMI). 2. 16% paraformaldehyde aqueous solution. 3. Permeabilization buffer: 1% Bovine serum albumin (BSA), 0.01% saponin, and Phosphate Buffered Saline (PBS). 4. Blocking solution: 1% BSA, 0.01% saponin, and PBS. (Optional: 1–5% serum, equal source as secondary antibody or 1–5% fetal bovine serum [FBS]). 5. STORM buffer: 50 mM Tris buffer (pH 8.0), 10 mM NaCl, 10% glucose, 0.1 M mercaptoethanolamine, 56 units/mL glucose oxidase, and 340 units/mL catalase. The STORM buffer is prepared by adding 100 μL of MEA solution and 10 μL of GLOX solution (see point 8 below ) to 890 μL of buffer B (from stock solutions listed below) to the sample just before imaging. 6. Buffer B: 50 mM Tris buffer (pH 8.0), 10 mM NaCl, 10% Glucose. 7. 1 M MEA: 77 mg of MEA (Cysteamine) in 1 mL of 0.25 N HCl (store at 4
C and prepare fresh every 15 days). 8. GLOX solution: 14 mg of glucose oxidase (from Aspergillus niger Type VII) and 1 mg catalase (from bovine liver) were dissolved in 250 μL of 10 mM Tris buffer (pH 8.0) 50 mM NaCl (store at 4
C and prepare fresh every 15 days). 9. Nikon stochastic optical reconstruction (STORM) microscope: For super-resolution imaging, we used a Nikon Ti STORM/ TIRF system equipped with four AOTF-controlled highpower lasers (405, 488, 561 and 647 nm), an Electron Multiplying Charge-Coupled Device (EMCCD) camera, and perfect focus system.

3

Methods

3.1 Isolation of Murine Neutrophils

1. Bone marrow cells are collected from the long bones of mouse legs by flashing the bones with 5 mL of PRF-RPMI using a 30 gauge ½00 hypodermic needle and a BD Luer-Lok™ 10-mL syringe.

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2. After flushing the bone marrow, the cells are collected in 10 mL of media in 15-mL Falcon tubes and spun down at 300  g for 10 min at 20
C. 3. During the isolation procedure, temperature fluctuation should be avoided to prevent possible cell activation. 4. The cells are then resuspended in PRF-RPMI by gentle pipetting using p200 pipettes and tips to help with bone marrow cell dispersion. 5. After addition of 10 mL RPMI, the cells are filtered using MACS SmartStrainers (30 μm), spun down, resuspended in 1 mL of isolation buffer, and isolated as follows. 6. There are several commercial and noncommercial methods available for the isolation of murine neutrophils, from Percoll gradient centrifugation [2] to positive and negative selection. We chose to use positive selection for highest purity using the Anti-Ly6G MicroBead Kit or Anti-Ly6G MicroBeads UltraPure. The latter has the advantage that it is a one-step methodology and thus is a faster method and utilizes a human antiLy6G monoclonal antibody, which does not interfere with most of the antibodies used for downstream applications. 7. Neutrophils are counted using a hemocytometer or a cell counter and then seeded on coverslips. We prefer a 4-Chamber 35-mm dish with 20-mm microwells. This dish has a #1.5 borosilicate bottom cover glass which is compatible with downstream applications including confocal, Airyscan, Total Internal Reflection Fluorescence Microscopy (TIRFM) and STORM. 3.2 Neutrophil Seeding

1. Add neutrophils (1–2  105/mL well) in 200 μL of PRF-RPMI (no serum) or HBSS. The choice of media is based on the downstream stimulation choice. 2. Optional: coat the glass with Poly-L-lysine. We incubate these wells with Poly-L-lysine at a concentration of 0.5 mg/mL in PBS w/o Ca2+ or Mg2+, 30 min to overnight at 37
C. If polyL-lysine is used, the medium must be completely removed, and the wells washed with PBS before adding polymorphonuclear leukocytes, (PMNs, neutrophils). 3. Incubate neutrophils for 1 h at 37
C in a tissue culture incubator under 5% CO2. 4. Remove the medium very carefully using a gel-loading tip with the pipette set up at 200 μL and with the tip in one of the corners of the well. It is important to be very careful at all steps before fixation to avoid removal of cells. 5. Optional: Wash the cells with PBS (w/o Ca2+ or Mg2+). Again, use a gel-loading tip to add and remove PBS. Add medium on

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the plastic corner of the well and let the media flow slowly into the bottom of the cover glass so as to avoid cell removal. Minimizing washes will prevent cells from detaching without necessarily increasing background. 3.3 Immunofluorescence Labeling

1. For immunofluorescence analysis, fix cells using 200 μL of 4% paraformaldehyde (PAF) (made fresh from 16%). Use gel-loading tips, and again, add the fixative on the plastic bottom of the well. Incubate for 8 min at room temperature (21
C). Remove PFA and wash with PBS w/o Ca2+ and Mg2+ (at least twice using 500 μL of washing solution). Once the cells are fixed, the slide can be inverted to remove media. Avoid vacuum systems. 2. Blocking: Incubate the cells with 1% BSA and 0.01% saponin in PBS (w/o Ca2+ and Mg2+) for 1 h at room temperature or overnight at 4
C. For some antibodies, to decrease nonspecific binding, blocking using 1–5% serum from the same source of your secondary antibody is recommended. Fetal bovine serum also works well for some antibodies. This should be determined empirically. 3. After blocking, remove the blocking solution and add the primary antibody in 1% BSA and 0.01% saponin in PBS (w/o Ca2+ and Mg2+). Saponin-mediated permeabilization is reversible, so it should be present in all incubations with primary and secondary antibodies. Incubate the primary antibodies overnight at 4
C with slight agitation. 4. Wash once with the blocking buffer and three times with PBS (500 μL, 5 min each). 5. Add the secondary antibodies in 1% BSA and 0.01% saponin in PBS (w/o Ca2+ and Mg2+). Incubate for 2 h at room temperature. 6. Wash once with the blocking buffer and three times with PBS as described above.

3.4

F-Actin Staining

1. The bicyclic heptapeptide, phalloidin binds to and stabilizes filamentous actin. To stain F-actin, add 5 μL of a 6.6 μM concentration of Alexa 488- or Alexa 647-Phalloidin in 200 μL of PBS with 0.05% Triton X-100. Remove media and add this solution to the cells. 2. Incubate for 30 min at room temperature. 3. Wash with 500 μL of PBS and with 1% BSA.

3.5

Storage

1. To store the samples, we use Fluoromount-G, which is a waterbased mounting media and thus does not interfere with and maintains the ideal refraction index required for downstream applications including TIRFM.

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2. When nuclei visualization is desired, 200 μL of FluoromountG-containing DAPI is added to the sample. Alternatively, DAPI can be added in the last wash, at a final concentration of 300 nM in PBS. Subsequently, the solution is removed, and the sample mounted in Fluoromount-G without DAPI. 3. Store the samples in the dark at 4
C until analysis. 3.6 Direct Stochastic Optical Reconstruction MicroscopeSuperResolution Microscopy

1. Super-resolutionmicroscopy using the Nikon STORM system is based on the stochastic optical reconstruction microscopy (STORM) technology that was developed by Xiaowei Zhuang’s laboratory at Harvard University [3]. In this method, the position of individual molecules is localized with high accuracy by switching them on and off sequentially using appropriate laser power settings and buffer conditions. 2. Cells seeded in glass bottom dishes (#1.5 borosilicate cover glass) are fixed with freshly made 4% paraformaldehyde for 10 min at room temperature, permeabilized, and blocked for 1 h with 1% BSA in PBS. The cells are labeled with appropriate primary antibodies in blocking solution for 2 h at room temperature, washed, and then incubated with the following secondary antibodies. 3. For two-color STORM, Alexa-647- and Atto-488 or Alexa488-conjugated secondary antibodies are used to label the proteins of interest. For three-color super-resolution imaging, Alexa-568-conjugated antibodies can be used as a third channel in addition to the above two sets of secondary antibodies (see Note 1). The samples are washed, post-fixed with freshly made 4% paraformaldehyde, and re-washed with water. Following this, either proceed to the next step or dry by inverting and store the samples in the dark for up to 48 h. 4. Prior to STORM imaging, the samples are incubated in freshly prepared STORM buffer, placed on the microscope stage, and then imaged using a 100  1.49 NA Apo TIRF objective either with or without TIRF illumination. 5. Images are collected at a frame rate of about 15 ms on 256  256 pixel region of the EMCCD camera using the multicolor sequential mode setting of the Nikon STORM module in Elements software. Three-dimensional (3D) STORM images are generated by introducing a cylindrical lens in the light path, which enables the assignment of Z position based on the shape of the point spread function [4]. Before STORM imaging, the objective is pre-calibrated for Z position assignment and for chromatic shift between the channels using 100-nm Tetraspeck beads as specified by Nikon. 6. After initially photo bleaching the samples using high laser power, the power on the lasers is adjusted, so that between

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50 and 300 molecules can be mapped per frame for each channel. Acquisition is stopped after a sufficient number of frames are collected (yielding 1–two million molecules), and the super-resolution images are reconstructed with the Nikon STORM software. 7. The positions determined from multiple switching cycles can show a substantial drift over the duration of the acquisition. The Nikon software automatically corrects for lateral and axial drift by using an autocorrelation method [5] wherein STORM images reconstructed from subsets of localizations at different time segments are aligned with those from the beginning of the acquisition to correct for drift. Axial drift over the course of the acquisition is also minimized by engaging the Nikon perfect focus system. 8. The position of well-separated molecules is estimated from the diffraction-limited images produced by the software and the precision of the localization during a switching cycle from the photon count of the individual switching event. Mean localization accuracy of about 20 nm is in good agreement with those reported previously, which can also be confirmed using Alexa488, -568, and -647 labeled IgG molecules that had been spread on coverslips at a low density. A Gaussian fit is used by the software to localize the position of each event to the final super-resolution image. 3.7 Stochastic Optical Reconstruction Microscopy Data Analysis for Protein– Protein Proximity

1. Images obtained on the Nikon N-storm (Nikon Inc) system are exported as localization coordinate map text files, which represent positions of individual blinks that have been localized with high accuracy by switching them on and off using the 488 nm, 561 nm, and 647 nm lasers. 2. In STORM, each fluorochrome can blink more than once and multiple blinks emanating from a molecule in the same 3D coordinate position defines the localization of that same molecule. The localization accuracy of this position is also recorded for each blink at each coordinate position. 3. The localization coordinates are imported into IMARIS software, where each blink is reconstructed as a sphere (spot) on an image grid; its centroid is the central coordinate position in three-dimensional space and the diameter of the sphere (spot) is the localization accuracy error (Fig. 1). 4. The widefield TIRF images are also imported into IMARIS as separate channels, aligned and fitted to overlay with the STORM image. This image is used to define cellular definition using the lesser resolved original fluorescent signals (Fig. 1). 5. The spots created are analyzed using the Colocalized Spots module to mark and score paired spots that lie within a defined

Fig. 1 Panels (a–g) pictorially display the process of analysis of TIRF and STORM data acquired on the Nikon TE2000, with representative images. Panel (a) shows a fluorescently labelled cell with actin (red) and MMP9 (green) that was imaged in TIRF microscopy mode. The fluorescent signals were auto-outlined in IMARIS using the isosurface module to generate a region of interest (ROI) outline as shown in panel (b). The STORM data were imported into this TIRF image file and registered as localization spots/points (spheres) as shown in (c). The data can also be displayed as coordinate positions without localization accuracy (sphere diameters) as shown in (d). In (e), we show the result of an analysis using the colocalization of spots module, displaying molecules that are 50 nm apart and superimposed over the original TIRF outlines. The regions magnified in (f) and (g) show that the colocalization image of spots was based on centroid position between two localized points, and the lateral localization is represented as the diameter of the spheres

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distance from each other in three-dimensional space. Specifically, the imported localization coordinate map of all fluorescent STORM confirmed blinks, which are previously filtered for drift and background signals in the NIKON software, are represented as spheres in IMARIS where the diameter of the sphere represents the localization accuracy and their centroid is used to compare distances between same and different paired molecules. 6. The Colocalized Spots module scores the number of each pair of fluorescent spots binned in the nanometer distance intervals as defined by the user. 3.8 Live Microscopy Analysis of Neutrophil Secretory Organelles and Actin Dynamics 3.8.1 Transfection of Primary Neutrophils

1. Bone marrow-derived neutrophils (BM-PMNs) are transfected by nucleofection using the Amaxa P3 Primary Cell 4D-Nucleofector X kit L and a 4D-Nucleofector System (Lonza AG, Allendale, NJ) [6]. 2. Briefly, 1–2.5  106 cells are resuspended in 20 μL of nucleofection solution P3 and are transferred to a nucleofection well. After addition of 2–3 μg of the expression vector of interest, the cells are immediately pulsed using the human monocytes setup in the 4D-Nucleofector System. 3. After the addition of 100 μL of PRF-RPMI medium (no serum), the cells are transferred to a 4-Chamber 35-mm dish with #1.5 borosilicate cover glass bottom. 4. The cells are recovered in a tissue culture incubator (5% CO2) and used in fluorescence-based assays 3–4 h after transfection. 5. Transfection efficiency using this method, depending on plasmid purity, is 20% [6]. Transfected cells are not preactivated by the nucleofection procedure and are responsive to stimulation.

3.9 Total Internal Reflection Fluorescence Microscopy

1. To study vesicle dynamics in live cells, cells transfected with appropriate vectors (GFP or mCherry conjugated to vesicle markers of interest) are resuspended in PRF-RPMI imaging medium (for neutrophils) or DMEM (for other cells) and seeded into a four-chamber 35-mm dish with #1.5 borosilicate cover glass bottom [7–9]. Cells are incubated at 37
C for at least 3 h before microscopy. 2. TIRF microscopy experiments are performed using a 100  1.45 numerical aperture TIRF objective on a Nikon TE2000-TIRF microscope equipped with a prewarmed microscope stage. 3. After placing the cells on the stage, the position of the individual laser beams is adjusted with the TIRF illuminator to impinge on the coverslip at an angle to yield a calculated evanescent field depth of a 100 nm (see Note 2).

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4. Images are acquired on a 14-bit, cooled charge-coupled device that is camera-controlled through NIS-Elements software. For live experiments, the images are recorded using 300–500-ms exposures depending on the fluorescence intensity of the sample. 5. For the quantitative analysis of the fluorescence intensity of vesicles in fixed cells, the exposure time and gain are kept unchanged throughout the experiment to enable comparative analysis of unstimulated and stimulated wild type and knockout cells. 6. Raw Nikon nd2 files are imported into IMARIS, where they are cropped in order to track and analyze all the vesicles in a single cell per field. The raw data are filtered using gaussian and median filters, and background-subtracted as well as autothresholded to ensure that signals are not missed due to mild photobleaching, thus allowing the use of all recorded frames acquired per time sequence. Vesicles are then tracked using surface rendering or spots modules in IMARIS. Both techniques were used and compared for accuracy of tracking vesicles and both techniques define vesicle displacement based on the position of the vesicle center or centroid. IMARIS software is used to calculate mean speed, acceleration, track length, and vector displacement. Data are imported into excel for further analysis and summary.

4

Notes 1. Since STORM imaging is carried out using a bandpass emission filter, care needs to be exercised in the choice of fluorophores used, so as to avoid issues with spillovers, especially when imaging three channels in the same sample. The intensity of staining in all three channels should be balanced to avoid using relatively high and low laser powers for different channels. Appropriate controls lacking one or both fluorophores may be used to check for signal due to spillover. 2. Depending on the cell type and the position of vesicles, minor iterative adjustments may be made to the TIRF angle and the Z position to make sure that the best image quality is obtained. The actual penetration depth can be recalculated for one of the color channels and the same calculated penetration depth can be set for other channels by using the formulae provided by Nikon. In this way, the same Z penetration depth is imaged for all channels.

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References 1. Ramadass M, Catz SD (2016) Molecular mechanisms regulating secretory organelles and endosomes in neutrophils and their implications for inflammation. Immunol Rev 273:249–265 2. Johnson JL, Ramadass M, Haimovich A, McGeough MD, Zhang J, Hoffman HM, Catz SD (2017) Increased neutrophil secretion induced by NLRP3 mutation links the Inflammasome to Azurophilic granule exocytosis. Front Cell Infect Microbiol 7:507 3. Rust MJ, Bates M, Zhuang X (2006) Subdiffraction-limit imaging by stochastic optical reconstruction microscopy (STORM). Nat Methods 3:793–795 4. Huang B, Jones SA, Brandenburg B, Zhuang X (2008) Whole-cell 3D STORM reveals interactions between cellular structures with nanometer-scale resolution. Nat Methods 5:1047–1052 5. Huang B, Wang W, Bates M, Zhuang X (2008) Three-dimensional super-resolution imaging by stochastic optical reconstruction microscopy. Science 319:810–813 6. Johnson JL, Monfregola J, Napolitano G, Kiosses WB, Catz SD (2012) Vesicular

trafficking through cortical actin during exocytosis is regulated by the Rab27a effector JFC1/ Slp1 and the RhoA-GTPase-activating protein Gem-interacting protein. Mol Biol Cell 23:1902–1916 7. Johnson JL, He J, Ramadass M, Pestonjamasp K, Kiosses WB, Zhang J, Catz SD (2016) Munc13-4 is a Rab11-binding protein that regulates Rab11-positive vesicle trafficking and docking at the plasma membrane. J Biol Chem 291:3423–3438 8. Zhang J, Johnson JL, He J, Napolitano G, Ramadass M, Rocca C, Kiosses WB, Bucci C, Xin Q, Gavathiotis E, Cuervo AM, Cherqui S, Catz SD (2017) Cystinosin, the small GTPase Rab11, and the Rab7 effector RILP regulate intracellular trafficking of the chaperonemediated autophagy receptor LAMP2A. J Biol Chem 292:10328–10346 9. Johnson JL, Hong H, Monfregola J, Kiosses WB, Catz SD (2011) Munc13-4 restricts motility of Rab27a-expressing vesicles to facilitate lipopolysaccharide-induced priming of exocytosis in neutrophils. J Biol Chem 286:5647–5656

Chapter 14 An Approach to Monitor Exocytosis in White Adipocytes Ali M. Komai, Man Mohan Shrestha, Saliha Musovic, and Charlotta S. Olofsson Abstract Exocytosis, the fusion of vesicles with the plasma membrane, can be measured with the patch-clamp technique as increases in membrane capacitance. Here we provide detailed information on how to monitor white adipocyte exocytosis using this method. We describe how to isolate the stromal vascular fraction of cells (SVF) within adipose tissue and how to differentiate SVF and cultured 3T3-L1 cells into adipocytes suitable for patch-clamp studies. We also give detailed protocols of how to record and analyze exocytosis in the differentiated cells. Key words Exocytosis, White adipocyte, Patch-clamp, Membrane capacitance, Whole-cell configuration, 3T3-L1 adipocytes, Stromal vascular cell fraction, Cell isolation, Adipocyte differentiation

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Introduction Historically, adipose tissue has been viewed as a rather inert organ, primarily important as a fat storage depot. However, growing evidence during the last two decades has led to the insight that white adipose tissue secretes a large variety of bioactive molecules, commonly referred to as adipokines, which have important roles in the control of whole body physiology [1]. In regard to mass, adipose tissue chiefly consists of large lipid-filled adipocytes. However, the white adipose tissue contains a number of other cell types. Those include macrophages, fibroblasts, vascular endothelial cells, and also preadipocytes, fibroblast-like cells that can form adipocytes during the process of adipogenesis. The non-adipocytes within adipose tissue are jointly referred to as the stromal vascular fraction (SVF; [2]). Adipokines are released both from the adipocyte itself and from SVF cells. A number of adipokines are, however, specifically secreted from the adipocyte. The real insight of the endocrine role of white adipose tissue came in the mid-1990s with the discovery of leptin [3] and

Florence Niedergang, Nicolas Vitale and Ste´phane Gasman (eds.), Exocytosis and Endocytosis: Methods and Protocols, Methods in Molecular Biology, vol. 2233, https://doi.org/10.1007/978-1-0716-1044-2_14, © Springer Science+Business Media, LLC, part of Springer Nature 2021

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adiponectin [4], two protein hormones secreted from the adipocyte itself. In fact, only adiponectin and leptin are clearly recognized as true adipose tissue-derived endocrine hormones defined by the fact that they are released from adipose tissue and exert distinct responses in target organs [1, 5]. Leptin has been shown to be secreted via vesicular exocytosis upon insulin stimulation, but its release also depends on Ca2+, although Ca2+ on its own is not sufficient to trigger leptin secretion [6, 7]; Ca2+ is instead required for insulin-mediated phosphorylation steps [7]. Already the first paper describing the discovery of adiponectin (initially termed Acrp30) suggested that this adipocyte hormone is released via regulated exocytosis [4]. However, until recently, the molecular control of its secretion has remained largely unknown. Work from our group has now begun to elucidate the mediators and mechanisms involved in the control of exocytosis of adiponectin. Vesicles containing adiponectin fuse with the adipocyte plasma membrane upon an elevation of Cyclic adenosine monophosphate (cAMP). Thus, similar to its role in leptin release, Ca2+ is important for augmentation of adiponectin secretion, but it does not act as a trigger of adiponectin exocytosis [8–11]. It is interesting that although adipocytes are clearly endocrine cells, the secretion of its two chief hormones is evidently not regulated in the same manner as in archetypical endocrine cell types where exocytosis is typically triggered by Ca2+ [12]. Adipocytes also secrete the protein adipsin (Factor D) upon exposure to insulin [13, 14]. Own results propose that insulin-stimulated adipsin secretion leads to an increase in plasma membrane capacitance, thus indicating vesicular exocytosis (Musovic et al., unpublished). Insulin moreover induces translocation of vesicles containing glucose transporter 4 (Glut4) to the adipocyte plasma membrane [15], to allow uptake of glucose. Vesicular exocytosis can be measured by the patch-clamp technique, as the increase in membrane capacitance occurs when vesicles fuse with the plasma membrane, thus enlarging its area [16]. Since their invention, capacitance recordings have been used to measure exocytosis in more or less every known endocrine and neuronal cell type. Capacitance measurements have also been applied to primary white adipocytes, but only in a couple of studies. In 1997, Lee and Pappone showed that extracellular ATP induced an ~16% increase of membrane capacitance in primary rat adipocytes [17]. A few years later, Robert Zorec et al. also showed a similar effect in adipocytes isolated from rat [18]. Whole-cell current recordings have also been achieved from primary adipocytes [19]. However, in all the three studies, recordings were performed at room temperature, perhaps to ease the sealing between the patch pipette and the plasma membrane, a procedure that is known to be more difficult at physiological temperatures. Performing membrane capacitance measurements at room temperature is problematic since regulated exocytosis is typically temperature-dependent.

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Own work has shown that this is also true for white adipocyte adiponectin exocytosis [9]. Altogether, several explanations can be found for the lack of electrophysiological studies of primary white adipocytes. One reason is of course the rather late discovery of the endocrine, hormone-secreting, function of white adipocytes. Another is the peculiar morphology of fat cells. Primary adipocytes are very large with a diameter of 20–150 μm in mice and 40–200 μm in humans [20]. This can be compared to other endocrine cell types that typically have a diameter of around 10 μm. Primary white adipocytes are packed with lipids and the triglyceride storage occupies ~95% of the cell and only allows a cytoplasmic rim of around 100 nm between the lipid and the plasma membrane (with exception for the cytoplasmic compartment closest to the nucleus). As a result, isolated primary adipocytes float (they are buoyanced by their large lipid content) and specific procedures are required in order to make them stick at the bottom of a cell culture dish and suitable for patch-clamp. Moreover, the white adipocyte plasma membrane is exceptionally rich with caveolae structures, invaginations that sometimes reach all the way to the unilocular lipid droplet [21]. The described specific characteristics of white adipocyte—their size, lipid content, and the largely folded plasma membrane—make those cells challenging for electrophysiological investigations. More importantly, reliable whole-cell recordings are difficult to achieve since the very narrow space between the plasma membrane and the triglyceride store obstructs electrical clamping of the entire (very large) cell. Primary adipocytes are also difficult to keep in culture, since they begin to lose much of their adipocyte features rather shortly after isolation [22]. Here we will describe a method to instead measure whole-cell membrane capacitance in adipocytes that have been differentiated in vitro from 3T3-L1 or SVF cells. Cultured adipocytes display several of the primary adipocyte functional characteristics and have, with regard to leptin [6, 23, 24] and adiponectin secretion, been demonstrated to be regulated in a similar way as primary adipocytes [9–11, 24]. 3T3-L1/SVF cells can easily be seeded and attached to the bottom of a dish and they stay attached even when they begin to accumulate lipid. Since they are not as lipidfilled as primary cells even when matured (their lipid is accumulated in the form of several smaller triglyceride stores and not as a single fat droplet), the ratio between cytoplasm and fat is larger and electrical contact can easily be achieved in the whole-cell configuration of the patch-clamp technique. In the next section, we will describe both the differentiation process and provide detailed information on how to measure exocytosis as an increase in membrane capacitance in cultured adipocytes.

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Materials All buffers and media are filtered using 0.2-μm filter units.

2.1 Isolation of Mouse Stromal Vascular Fraction Cells

1. Krebs-Ringer Glucose HEPES (KRGH) buffer: 25 mM HEPES, pH 7.4, 1.2 mM KH2PO4, 118 mM NaCl, 4.7 mM KCl, 1 mM MgSO4, 1 mM CaCl2, and 5.5 mM dextrose. 2. Wash buffer: KRGH buffer with 3% bovine serum albumin (BSA). 3. Digestion buffer: KRGH buffer with 1.5% BSA and 1 mg/ml collagenase type II. 4. Stromal vascular fraction (SVF) medium: High-glucose Dulbecco’s modified eagle medium (4500 mg/L glucose DMEM) with 10% fetal bovine serum (FBS, Hyclone) and 1% penicillin/ streptomycin (P/S). 5. 70% Ethanol. 6. Styrofoam, needles, forceps, scissors, scale, 37
C shaker, 100-μm sterile cell strainer, 30-μm sterile cell strainer, centrifuge, syringe with long needle, serological pipettes, pipette-aid, collagen-coated 35-mm tissue culture dishes.

2.2 Culture and Differentiation of Mouse Stromal Vascular Fraction Cells

1. Phosphate-buffered saline (PBS): 37 mM NaCl, 10 mM Na2HPO4, 2.7 mM KCl, 2 mM KH2PO4, and pH 7.4. 2. Stromal vascular fraction (SVF) medium: High-glucose Dulbecco’s modified eagle medium (4500 mg/L glucose DMEM) with 10% FBS and 1% penicillin/streptomycin (P/S). 3. Stromal vascular fraction adipogenic cocktail: SVF medium supplemented with 850 nM insulin (human recombinant; Actrapid Penfill), 1 μM dexamethasone (DEXA), 0.5 mM 3-isobutyl-1-methylxantine (IBMX), and 1 μM rosiglitazone. 4. 1 mM dexamethasone (1000-fold concentration) dissolved in water or ethanol depending on its formulation. 5. A 10 mM stock solution of rosiglitazone (10,000-fold concentration). 6. A 500 mM stock solution of IBMX (1000-fold concentration) dissolved in 50 mM NaOH. 7. A 100 IU/ml (~596 μM) of insulin. 8. Trypsin, EDTA. 9. 35-mm tissue culture dish, 37
C incubator, centrifuge, 37
C water bath, serological pipette, and pipette aid.

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1. 3T3-L1 preadipocytes (passage 9–15)—original vial from Zen-Bio (#SP-L1-F). 2. Proliferation medium: high-glucose Dulbecco’s modified eagle medium (4500 mg/L glucose DMEM) containing 10% New Born Calf Serum (NBCS, Life technologies), and 1% penicillin–streptomycin (P/S). 3. Differentiation medium 1 (D1): high-glucose DMEM, 10% fetal bovine serum (FBS), 1% P/S, 850 nM insulin (human recombinant; Actrapid Penfill), 1 μM dexamethasone (DEXA), and 0.5 mM IBMX. 4. Differentiation medium 2 (D2): high-glucose DMEM, 10% FBS, 1% P/S, and 850 nM insulin. 5. Trypsin. 6. T75 cell culture flasks, 35-mm tissue culture dishes, 37
C incubator, centrifuge, 37
C water bath, serological pipette, and pipette-aid.

2.4 Electrophysiological Measurements

1. Equipment: Anti-vibration table with mounted Faraday cage, inverted microscope with 20–30 air objective, patch-clamp amplifier (HEKA Electronics, Germany), headstage, macromanipulator, micromanipulator, computer, recording software (PatchMaster), peristaltic perfusion pump, temperature controller, Osmometer, pipette puller, horizontal pipette holder with tip heater, pipette polisher (Microforge or other), pipette box (covered with a lid for storing and protection of prepared pipettes). 2. Consumables: borosilicate glass capillaries (O.D 1.5 mm, I.D 1.17 mm), Sylgard (Dow Corning, USA), perfusion tubing, culture dish patch-clamp insert (see Fig. 5a). 3. Extracellular (EC) solution for perfusion of the cell culture dish. Recommended basic EC solution (see Note 1): 140 mM NaCl, 3.6 mM KCl, 2 mM NaHCO3, 0.5 mM NaH2PO4, 0.5 mM MgSO4, 5 mM HEPES (pH 7.4 with NaOH), 2.6 mM CaCl2, and 5 mM glucose. Prepare solution using mQ-water. Store EC solutions refrigerated (see Note 2). 4. Recommended basic IC solution (see Note 1): 125 mM K-glutamate, 10 mM KCl, 10 mM NaCl, 1 mM MgCl2, and 5 mM HEPES (pH 7.15 with KOH). Aliquot and store IC solutions at 20
C (typically in 1-ml portions).

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Methods

3.1 Isolation of Mouse Stromal Vascular Fraction Cells

Mouse adipose stromal vascular fraction is isolated as previously described [25] with some modifications. A schematic illustration of the steps involved in isolation of mouse SVF cells from mouse white adipose tissue (WAT) is shown in Fig. 1. 1. Sacrifice the mouse using anesthesia or by CO2 asphyxiation as approved by your local guidelines on euthanasia. Saturate the fur with 70% ethanol and place the mouse with its ventral side up and fix it on Styrofoam with needles. After euthanizing, flush the mouse properly with 70% ethanol to avoid fur getting into the tissue extracts and contaminating the cells. 2. Pull up the abdominal skin using sterile forceps and cut the skin vertically toward the head and along the lower limbs, the cut should look like an inverted “Y.” 3. Open the abdominal cavity and remove the desired white fat depots using sterile forceps and scissors. 4. Weigh the tissue and place it in 50-ml conical tube containing 5 ml per mouse wash buffer. 5. Transfer the rinsed tissue to a new 50-ml conical tube and mince the tissue thoroughly into small pieces using sterile scissors. 6. Add 5 ml digestion buffer per mouse and shake at 120 rpm in 37
C for 45–60 min until the tissue appears smooth on visual inspection. Always use freshly prepared digestion buffer to maintain the activity of the collagenase. 7. After collagenase digestion, pipette the solution up and down ten times using a 10-ml pipette and filter the solution through a sterile 100-μm cell strainer to remove undigested tissues. Be careful not to produce bubbles while pipetting. 8. Add 15 ml of SVF medium to dilute the digestion buffer and centrifuge at 600  g for 5 min to pellet the stromal vascular cells. 9. Aspirate the floating lipid and adipocyte layers using a syringe with long needle. Aspirate from the top, so that the lipid layer is removed as much as possible. 10. Resuspend the stromal vascular cells in 5 ml per mouse SVF medium. Avoid producing bubbles while resuspending the cells. 11. Filter the solution using 30-μm cell strainer and dilute the flow through in 15 ml per mouse SVF medium. 12. Centrifuge the resuspended cells at 600  g for 5 min and carefully aspirate the supernatant.

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Fig. 1 Schematic illustration of SVF cell isolation from mouse WAT. The illustration was made using “Affinity Designer” software

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13. Resuspend the cells in 2.5 ml per mouse SVF medium and plate 2 ml of cell suspension onto collagen-coated 35-mm tissue culture dishes or glass cover slips (see Note 3). 3.2 Proliferation of 3T3-L1 Adipocytes

1. Take up a vial of 3T3-L1 preadipocytes (300,000–500,000 cells/vial) from liquid nitrogen and place in a 37
C water bath and leave until nearly thawed (only a small piece of ice visible). 2. Immediately add cells to a sterile 15-ml falcon tube containing 10 ml of proliferation medium. Centrifuge at 200  g at room temperature (RT) for 4 min. Aspirate the medium and resuspend cells in 10 ml volume of proliferation medium. 3. Place the cell suspension in a T75 cell culture flask and keep in a humidified incubator at 37
C and 5% CO2. 4. Maintain the cells in the flask until they reach 60% confluency (~2 days). 5. Wash the cells twice with PBS and add 1.5 ml of 0.25% trypsin and incubate for 3 min at 37
C. 6. Upon trypsin treatment, resuspend the cells in 10 ml of proliferation medium and centrifuge at 200  g at RT for 4 min. 7. Remove medium and resuspend cell pellet with 10 ml of fresh proliferation medium and seed on 35-mm tissue culture dishes or glass cover slips (see Note 3). 8. Proliferate cells for another 2 days until they reach 90% confluency (see Note 4).

3.3 Differentiation of Stromal Vascular Fraction Cells

In vitro differentiation is performed as previously described [25] with some modifications. The adipogenic differentiation of SVF cells is shown in Fig. 2. 1. Incubate the plated cells in 37
C incubator in the presence of 5% CO2. Always use a clean and sterile primary cell culture incubator to avoid contaminations. 2. Twenty-four hours after plating, aspirate the medium and rinse the cells with PBS pre-warmed to 37
C and aspirate. Check the cells under an inverted microscope to make sure that the unattached cells are removed; if the unattached cells remain after the first rinse, repeat the rinse with PBS and aspirate. Rinsing the cells should be done quickly to avoid the cells from drying out. 3. Add fresh SVF medium. 4. Change to fresh SVF medium every 48 h until the cells are 80–90% confluent (see Note 5). 5. Once the cells reach 80–90% confluence, aspirate the medium and rinse the cells once with pre-warmed PBS and add 0.5 ml of 0.25% trypsin.

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Fig. 2 In vitro adipogenic differentiation of SVF cells and Oil Red O staining of mature adipocytes. Scale bar ¼ 500 μm, magnifications are 10 and 20

6. Incubate for 3 min at 37
C and add 5 ml SVF medium to dilute the trypsin. 7. Centrifuge at 600  g for 5 min and aspirate the supernatant. 8. Resuspend the cells in SVF medium with 1:3 to 1:5 dilution and plate 2 ml of cell suspension onto collagen-coated 35-mm tissue culture dish/glass cover slip. Culture the cells until 80–90% confluence (see Note 5). 9. Forty-eight hours after reaching 80–90% confluence, aspirate the medium and rinse the cells twice with pre-warmed PBS. 10. Replace the SVF medium with the SVF adipogenic cocktail and mark the culture as Day 0 of differentiation.

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Fig. 3 In vitro differentiation of 3T3-L1 adipocytes. Differentiation from fibroblast like cells (day 0) to mature adipocytes expressing multiple lipid droplets in the cytoplasm (day 8–9). Scale bar: 50 μm

11. After 48 h, replace the SVF adipogenic cocktail with SVF medium supplemented with 850 nM insulin. 12. Aspirate the medium and replace it with fresh SVF medium supplemented with 850 nM insulin every 48 h until harvest when cells are visually differentiated (see Note 6). 3.4 Differentiation of 3T3-L1 Adipocytes

The changes of cell density and morphology during the stages of differentiation are shown in Fig. 3. 1. Remove the proliferation medium. 2. Initiate differentiation of 3T3-L1 adipocytes with differentiation medium D1. 3. After 2 days, remove differentiation medium D1 and replace it with fresh differentiation medium D2. 4. Add fresh proliferation medium every 48 h up until day 8–9 after the initial start of differentiation, when cells are visually differentiated (see Note 7). 5. Performed measurements of exocytosis on day 8 or 9 (see Note 8).

3.5 Preparation of Patch Pipettes

Prepare fresh pipettes every day. A patch pipette used for capacitance measurements is produced in three steps: 1. By use of a pipette puller, make borosilicate glass pipettes with a resistance between 2.5 MΩ and 3.5 MΩ when filled with the intracellular solution (a higher pipette resistance will make whole-cell recordings more difficult). There are different types of pipette pullers available and heater settings need to be adjusted in agreement with manufacturer’s recommendations. Settings typically need to be calibrated until pipette tips are satisfactory. 2. Use a horizontal pipette holder with tip heater and place the pipette so that the tip is visible in a light microscope. While viewing the tip, use a bent syringe needle to carefully coat the

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outside of the tip with Sylgard (prepared according to manufacturer’s protocol) while taking care not to clog the pipette tip opening. While manually turning the pipette, heat the tip for ~3 s or until the Sylgard is solid. The Sylgard should cover the part of the pipette that will dip into the perfusion solution during the capacitance recording. It is critically important to coat the pipette for measurements of exocytosis. The thickening/isolation of the glass reduces the capacitance between the glass walls and the bath; this capacitance can otherwise disturb the recordings. 3. Fire-polish the edges of the pipette tip by heating by use of a pipette Microforge; note that this process will decrease the size and alter the shape of the pipette tip (the magnitude of changes depend on heating temperature and duration). Adjust settings to achieve a pipette tip with the final resistance of 2.5–3.5 MΩ. Figure 4 shows the process of coating a pipette tip with Sylgard viewed through the ocular (A–C), the in-house-produced horizontal pipette holder with tip heater (D), and the pipette storage box (E). 3.6 Patching Adipocytes: Attaining a GΩ Seal and Electrical Contact

Although this protocol provides detailed information on how to carry out electrophysiological measurements of membrane capacitance in white adipocytes, it is highly recommended that the experimenter is experienced in electrophysiological measurements applying the patch-clamp technique. Please note that some specific adjustments may be required when patching adipocytes compared to more conventional cell types (see Note 9). 1. Start the patch-clamp amplifier, the computer and patch-clamp software and turn on the microscope. 2. Take out a dish of cells from the incubator, remove medium and wash twice with EC solution using a 1-ml pipette (do this gently in order not to rinse away the cells!). Leave a small volume of solution in the dish to not let cells dry out before mounting on the patch setup. 3. Mount the cell culture dish on the patch-clamp setup and carefully place the dish insert into the dish. Secure the dish and the insert with adhesives such as Blu Tack as to prevent movement and vibrations. Several alternatives are available for the recording chamber, and cells may be cultured on glass cover slips and mounted, depending on the setup. Figure 5a shows the in-house-produced insert, fitting into a 35-mm cell culture dish. 4. Insert both perfusion tubes (inlet and outlet) and the reference electrode into the dish/recording chamber. Make sure the reference electrode is in contact with the solution perfusing the dish in order to obtain an electric circuit.

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Fig. 4 Preparation of Patch pipettes. A pipette (a) together with a bent syringe tip dipped in Sylgard (b) and a pipette with Sylgard coating(c). (d) shows an inhouse-produced horizontal pipette holder with tip heater and e an in-houseadjusted pipette storage box

Fig. 5 Patch-clamp insert and mounted cell dish. In-house-produced insert made of plexiglass (a) and insert fitted to a mounted cell dish with reference electrode, perfusion inlet and outlet, and a patch pipette containing IC solution (b)

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5. Start the perfusion and temperature controller and perfuse the cells for ~5–10 min before attempting experiments. 6. Make sure that temperature controller settings are adjusted to allow desired temperature at the center of the dish (this might require insulation of the inflow tubing as to prevent the solution to lose heat on its way through the tubing to the dish). It is critically important to maintain a near physiological and stable temperature during the recordings (see Note 10). Figure 5b shows a mounted cell dish ready to patch. 7. Find a lipid-filled cell (some cells might still be more fibroblast looking), preferably with no lipid accumulation at the location where the pipette tip will be attached (see Note 11). 8. Fill a micropipette with IC solution to ~1/4 of the pipettes length using a 1-ml syringe. Carefully flick/tap the tip to release small air bubbles. When no more bubbles can be observed after flicking, mount the pipette on the intracellular electrode. 9. Adjust the patch-clamp headstage with pipette to a desired angle (an angle of ~45
is recommended to facilitate pipette sealing), with the pipette tip ready to enter the dish. 10. Use the macromanipulator to lower the pipette tip close to the solution, just above the fluid level. 11. Apply light positive pressure to the suction tube and lower the pipette to enter the solution/dish—be careful not to lower the pipette too fast as to prevent the pipette tip from hitting the bottom and breaking. You will know that you have entered the solution by monitoring the resistance on the computer screen. Correct the pipette offset caused by the liquid junction potential. 12. Lower the pipette with small movements using the macromanipulator and make sure to keep focus on the tip for each step, until the pipette tip is just above the cell you aim to patch. 13. Now continue to lower the pipette carefully using the micromanipulator, until the pipette tip touches the membrane and its resistance is approximately doubled. Clamp the cell stepwise at negative voltages to facilitate sealing. Finally, clamp the membrane potential at 70 mV (to ascertain that voltagedependent currents do not disturb the capacitance recording). 14. Apply a steady gentle suction/negative pressure to gain the GΩ seal. If the resistance increase is very slow or halts, continue to keep the steady suction/negative pressure for ~10–20 s. However, should the seal fail within this time, stop and repeat from step 7. The likelihood of obtaining a seal is low when the first attempt fails and it is in our experience better to start over with a new cell and a new pipette.

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Fig. 6 Graphical representation of the patch-clamp procedure. The process of attaining a GΩ seal, as well as gaining access to the cell interior by rupturing of the plasma membrane and thus obtaining the standard whole-cell configuration

15. When a GΩ seal has been attained, apply pulses of gentle suction stepwise, increasing the force until the membrane patch between the pipette and the cell interior is ruptured and electrical contact is established. It is necessary to be gentle during this process, in order not to lose the GΩ seal. Figure 6 shows the procedure of attaining a GΩ seal and electrical access by rupture of the plasma membrane. 16. Monitor access resistance and the leak current during the experiment. The access resistance should be stable and below ~15 MΩ. If resistance is unstable or increases, again apply gentle suction to regain satisfactory access. Should the leak current increase, try applying a steady gentle suction for a short duration of time to attempt improving the seal. Figure 7a illustrates a simplified representation of a patchclamp circuit in the standard whole-cell configuration. Figure 7b shows an adipocyte with attached patch pipette. 3.7 Membrane Capacitance Recordings

1. Make sure to compensate for capacitive currents over the patch pipette before a measurement is initiated. Start your capacitance recording. The recording protocol depends on the used software. Here we use PatchMaster software (HEKA, Germany) and register exocytosis by repetitive Auto C-slow compensation using the Cap Track function. This procedure applies short trains of square-wave pulses and averages the resulting currents and fit and exponential to deduce the compensation values necessary to cancel the current. The number of currents and their amplitude can be varied—we typically apply a train of 10 pulses with an amplitude of 5 mV. The recording may proceed over several minutes and a stable seal and access can sometimes be maintained 30 min. Should the access resistance increase during the recording, try to apply gentle suction to regain satisfactory access. Should the leak current increase, try applying a steady gentle suction for a

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Fig. 7 (a) Simplified representation of a patch-clamp circuit in the standard whole-cell configuration. After gaining the seal, the disruption of the membrane allows wash-in of the pipette solution and completion of an electrical circuit between the pipette and reference electrode. The voltage clamp mode allows the cell to be clamped at a specific membrane potential to monitor the membrane capacitance (Cm). Rm represents the membrane resistance (b) A 3T3-L1 adipocyte 8 days after start of differentiation with attached patch pipette. Scale bar ¼ 50 μm

short duration of time to improve the seal. Note carefully that the suction does not affect the recorded capacitance trace. 2. Stimulation of exocytosis with the IC (pipette) solution: Start recording the capacitance rapidly after achieving a steady seal and access, in order to enable recording of a stable part of membrane capacitance (before exocytosis is triggered by rinsing in of the intracellular solution; see Note 12). Typically, endocrine cell exocytosis initially proceeds at a fast rate that ceases with time (depending on the exocytosis of vesicles belonging to different functional pools with different release characteristics [12, 26]). Record the capacitance for as long as possible or until you have been able to observe the desired exocytosis response.

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3. Stimulation of exocytosis via the EC solution: Start recording the capacitance when a steady seal and access has been attained. Record the capacitance at 1 or 2 min in order to have a part of the capacitance trace where no changes are observed (see Note 12). Add the secretagogue (a compound, agent, or hormone expected to trigger exocytosis) to the perfusion (EC) solution rinsing into the dish. Take a note of when the solution containing the secretagogue enters the cell culture dish (see Note 13). Continue the recording during the time required to observe the expected effect. Figure 8a shows a typical capacitance recording in a 3T3-L1 adipocyte and indicates the triggering of exocytosis via IC or EC solution. For some general patch-clamp advice, see Note 14. 3.8 Analysis of Membrane Capacitance Recordings

4

The rate of exocytosis (capacitance increase) can be measured at one or several time-points by application of linear fits using OriginPro (OriginLab Corporation) or another data analysis software. The total capacitance increase at the end of the recording, reflecting the number of vesicles released, as well as the duration of exocytosis (if a plateau was reached when net exocytosis is no longer measurable) may also be determined. Examples of analyses are illustrated in Fig. 8b. The rate of capacitance increase is typically expressed and plotted as femtofarad/second (fF/s). If relevant, the analysis of exocytosis may be expressed in relation to cell size (see Note 12). For further examples of how to carry out, analyze, and present adipocyte membrane capacitance recordings, see [9–11].

Notes 1. As a general recommendation, the aim should be to prepare solutions that are close to physiological conditions with regard to concentrations of salts and pH. Mg-ATP, cAMP, and/or Ca2+ (mediators involved in regulation of vesicle exocytosis) may be included at chosen concentrations. The Ca2+chelators, EGTA or BAPTA, can be included to buffer intracellular Ca2+ at appropriate free concentrations (the free [Ca2+] of IC solutions can be calculated using Maxchelator, Stanford). 2. The EC solution can be used for up to 1 week, but make sure that no contamination is visible. Bring the solution to room temperature before starting your experiment—the temperature of the heated EC solution perfusing the cell dish may vary depending on its initial temperature. 3. Cells may be cultured on differently sized coated dishes or glass cover slips, depending on what fits the experimental setup. Here we use 35-mm plastic dishes where cells are mainly seeded

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Fig. 8 (a) An example of a typical capacitance measurement. To stimulate exocytosis, the cell can be infused with and IC solution containing a triggering mediator/compound (IC stim). Alternatively, the adipocyte may be infused with a non-stimulatory pipette solution and exocytosis triggered after addition of a secretagogue-containing EC solution, perfusing the dish (EC stim). Note that infusion with a stimulatory pipette solution often results in the rapid onset of exocytosis shortly after gaining access to the cell (not represented in the illustration); it is thus important to quickly start the recording. (b) Illustration of how to analyze capacitance recordings. Exocytotic rates (ΔCm/Δt; fF/s) can be determined by application of linear fits (red) to the capacitance trace at different time points, as indicated by the vertical dotted lines. The total capacitance increase, reflecting the total amount of exocytosis/released vesicles, is measured on the y-axis from the initial capacitance value until the point where exocytosis ceases and a plateau is reached. The duration or exocytosis is measured from the start of capacitance increase until a plateau is reached

in the middle (to avoid usage of an unnecessarily large number of cells). When patching the cells, you may be unable to reach cells grown closer to the edges of the dish. 4. If the desired confluency to proceed to differentiation is not achieved within 2 days, you may grow the cells longer. In that case, add fresh proliferation medium every 48 h. 5. This typically takes 2 days. However, the cells may require different number of days in order to reach the right degree of confluency.

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6. Typically, the cells are well differentiated on day 8 but more time may be required. The differentiation degree can be visually judged by the accumulation of lipid droplets. 7. The degree of differentiation can be visually judged by lipid droplet accumulation in the cytoplasm. 8. Cells can be used for electrophysiological experiments at day 7–9 from start of differentiation. At later days, [9, 10] adipocytes are often more difficult to patch (complicated to get and keep a seal), likely due to the increased lipid content. 9. Even as an experienced electrophysiologist, you may find that patching adipocytes is a challenge. Due to the large lipid content of those cells and abundant caveolae structures, the adipocyte plasma membrane is different from that of conventional cell types. Greater care may need to be taken when applying suction in order to achieve a GΩ seal (a robust contact between the patch pipette and the cell membrane) and when attaining the standard whole-cell configuration (rupture of the plasma membrane located in the tip of the patch pipette). Due to the lipid filling, it may also be more difficult to see your patching process in the inverted microscope, especially if cells are densely seeded. A large portion of patience and determination is a must. 10. Exocytosis in neuronal and endocrine cell types is typically temperature-dependent [27, 28]. This is true also for the white adipocyte exocytosis of adiponectin [9]. However, it is recommended not to go above 32–33
C since it is usually very difficult to achieve good contact between the patch pipette and the cell membrane (a so called GΩ seal) at higher temperatures. 11. The occurrence of lipid at the spot where the patch pipette is attached easily results in the lipid entering the pipette tip and clogging it. Although this can sometimes be removed by applying gentle positive pressure, this is not advised since the seal often breaks during this procedure. Moreover, a small amount of lipid often stays at the tip, something that can alter the pipette resistance. If exocytosis is stimulated by the IC solution in the patch pipette, applying positive pressure may also result in premature triggering of exocytosis (before being able to start the recording). 12. It is important to be aware of the fact that recordings of membrane capacitance measure the net balance between endocytosis and exocytosis. The IC solution is typically composed so as to minimize endocytosis, but endocytosis may still occur during the experiment. Likewise, sometimes a small magnitude of exocytosis can be observed before application of the stimulus. To ascertain that the recorded change of membrane capacitance is truly due to the applied stimulus, it is helpful to be able

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to analyze the initial stable rate (close to 0). Experiments where cells at the start of the recording display a significant capacitance change (up or down) should be discarded. Moreover, it may be useful to also record the initial cell size; this can be utilized when capacitance responses are analyzed. 13. It is important to be aware of the fact that the wash-in of a secretagogue via the perfusion solution does not rapidly and immediately expose the cell to the absolute concentration in the solution—a gradual change over time will occur until the solution in the dish/chamber is exchanged. Fast perfusion systems are available for a more rapid application. The studied agent/compound may also be applied in close vicinity to the patched cell by local puff application [29]. 14. The patch-clamp method is overall a demanding technique and obstacles in the form of pipette drift or “noise” disturbing the recordings may be experienced. Some general advices are to exclude pipette drift (leading to lost GΩ seals) by attaching a pipette to the headstage, mark a small spot at the bottom of an empty dish, fill the dish with solution, place the pipette tip just above the mark and wait ~15 min. When you view the pipette again, you will see if its position has changed. In case of drift while using a hydraulic micromanipulator with water, it might need to be refilled. Another common cause of drift is instability of the headstage cable.

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Concluding Remarks Although the application of capacitance recordings to adipocytes might be challenging, more studies of the kind described here are needed in order to understand the endocrine/secreting role of white adipocytes. The method outlined here may of course be applied to other in vitro differentiated adipocytes, for example, of human origin (make sure to adapt the proliferation and differentiation protocols to fit the cell type). While membrane capacitance recordings yield detailed information at the single cell level, the method does unfortunately not measure what is actually secreted. The exocytosis data can be compared to secreted product from cells exposed to similar conditions during shorter time periods (incubation time chosen to match the duration of a capacitance recording), as has been done with regard to adiponectin exocytosis [8–11]. However, in order to directly study exocytosis of a specific vesicle type, advanced imaging techniques must be used. Total internal reflection fluorescence microscopy (TIRFM) applied to live cells is one of the most powerful imaging techniques that delivers necessary spatiotemporal resolution in order to image intracellular processes, such as adipocyte vesicle dynamics and

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exocytosis [30]. Cells may be transfected in order to express the fluorescently labelled protein of interest (leptin, adiponectin, or other adipokine). Direct visualization of adipokine vesicles using TIRFM would advance the understanding of the molecular regulation of exocytosis in the white adipocyte and greatly progress the understanding of the endocrine role of this cell type.

Acknowledgments This work was supported by the Swedish Medical Research Council (grant 2013-7107). We thank Mr. Habz Nafar for the illustration in Fig. 7a. References 1. Ghaben AL, Scherer PE (2019) Nat Rev Mol Cell Biol 20:242–258 2. Trujillo ME, Scherer PE (2006) Endocr Rev 27:762–778 3. Zhang Y, Proenca R, Maffei M, Barone M, Leopold L, Friedman JM (1994) Nature 372:425–432 4. Scherer PE, Williams S, Fogliano M, Baldini G, Lodish HF (1995) J Biol Chem 270:26746–26749 5. Scheja L, Heeren J (2019) Nat Rev Endocrinol 15:507–524 6. Wang Y, Ali Y, Lim CY, Hong W, Pang ZP, Han W (2014) Biochem J 458:491–498 7. Cammisotto PG, Bukowiecki LJ (2004) Am J Physiol Regul Integr Comp Physiol 287: R1380–R1386 8. Brannmark C, Lovfors W, Komai AM, Axelsson T, El Hachmane MF, et al. 2017. J Biol Chem 9. El Hachmane MF, Komai AM, Olofsson CS (2015) PLoS One 10:e0119530 10. Komai AM, Brannmark C, Musovic S, Olofsson CS (2014) J Physiol 592:5169–5186 11. Komai AM, Musovic S, Peris E, Alrifaiy A, El Hachmane MF et al (2016) Diabetes 65:3301–3313 12. Burgoyne RD, Morgan A (2003) Physiol Rev 83:581–632 13. Cook KS, Min HY, Johnson D, Chaplinsky RJ, Flier JS et al (1987) Science 237:402–405 14. Lo JC, Ljubicic S, Leibiger B, Kern M, Leibiger IB et al (2014) Cell 158:41–53 15. Stockli J, Fazakerley DJ, James DE (2011) J Cell Sci 124:4147–4159

16. Lindau M, Neher E (1988) Pflugers Arch 411:137–146 17. Lee SC, Pappone PA (1997) Pflugers Arch 434:422–428 18. Chowdhury HH, Grilc S, Zorec R (2005) Ann N Y Acad Sci 1048:281–286 19. Zhang Y, Xie L, Gunasekar SK, Tong D, Mishra A et al (2017) Nat Cell Biol 19:504–517 20. Hagberg CE, Li Q, Kutschke M, Bhowmick D, Kiss E et al (2018) Cell Rep 24:2746–56 e5 21. Thorn H, Stenkula KG, Karlsson M, Ortegren U, Nystrom FH et al (2003) Mol Biol Cell 14:3967–3976 22. Jumabay M, Bostrom KI (2015) World J Stem Cells 7:1202–1214 23. Zeigerer A, Rodeheffer MS, McGraw TE, Friedman JM (2008) Exp Cell Res 314:2249–2256 24. Xie L, O’Reilly CP, Chapes SK, Mora S (2008) Biochim Biophys Acta 1782:99–108 25. Shao M, Vishvanath L, Busbuso NC, Hepler C, Shan B et al (2018) Nat Commun 9:890 26. Rorsman P, Renstrom E (2003) Diabetologia 46:1029–1045 27. Bittner MA, Holz RW (1992) J Biol Chem 267:16226–16229 28. Renstrom E, Eliasson L, Bokvist K, Rorsman P (1996) J Physiol 494(Pt 1):41–52 29. Wu Z, Auclair SM, Bello O, Vennekate W, Dudzinski NR et al (2016) Sci Rep 6:27287 30. Nagamatsu S, Ohara-Imaizumi M (2008) Methods Mol Biol 440:259–268

Chapter 15 Amperometry in Single Cells and Tissue Damien J. Keating Abstract The release from cells of signaling molecules through the controlled process of exocytosis involves multiple coordinated steps and is essential for the proper control of a multitude of biological pathways across the endocrine and nervous systems. However, these events are minute both temporally and in terms of the minute amounts of neurotransmitters, hormones, growth factors, and peptides released from single vesicles during exocytosis. It is therefore difficult to measure the kinetics of single exocytosis events in real time. One noninvasive method of measuring the release of molecules from cells is carbon-fiber amperometry. In this chapter, we will describe how we undertake such measurements from both single cells and in live tissue, how the subsequent data are analyzed, and how we interpret these results in terms of their relevant physiology. Key words Exocytosis, Amperometry, Chromaffin cells, Enterochromaffin cells, Serotonin, Catecholamines, Adrenaline, Noradrenaline

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Introduction Regulated exocytosis has been studied over many years in a number of model systems, using an array of both in vivo and in vitro approaches. Amperometry is a method that has been used since the early 1970s to study exocytosis of neurotransmitters and hormones [1]. It takes advantage of the fact that a suitable potential can oxidize some chemicals, such as amines, resulting in a measurable flow of current. It is these changes in current upon the oxidation of such molecules that represent the measurable components that are studied using amperometry, and which provide valuable information regarding the mechanisms controlling this coordinated cell secretion. Amperometry was originally used in the biological sciences to measure secretion in whole-tissue samples, in particular, looking at neurotransmitter release in the brain [1, 2]. A number of the molecules, which undergo exocytosis, are capable of being oxidized, including amines such as noradrenaline (norepinephrine), adrenaline (epinephrine), dopamine and

Florence Niedergang, Nicolas Vitale and Ste´phane Gasman (eds.), Exocytosis and Endocytosis: Methods and Protocols, Methods in Molecular Biology, vol. 2233, https://doi.org/10.1007/978-1-0716-1044-2_15, © Springer Science+Business Media, LLC, part of Springer Nature 2021

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serotonin. The technique of amperometry utilizes a polarizable electrode in solution, to which a voltage greater than the redox potential of the species in question is applied [3]. In the presence of oxidizable chemicals released from vesicles during exocytosis, molecules diffusing close to the electrode surface are oxidized, transferring electrons to the electrode, and resulting in current flow that is measured using circuitry that operates similar to a patch-clamp amplifier in voltage-clamp mode. This chapter will discuss the use of amperometry in which the voltage applied to the electrode is held constant and how this is used to measure exocytosis in real time from cells and tissue. While providing significant information on the release of neurotransmitters within the brain in the in vivo situation, the full power of amperometry to investigate exocytosis was not appreciated until these techniques were extended to the single-cell level [4]. By placing the electrode in close vicinity of a single cell, exocytosis can be measured with sufficient temporal and spatial resolution to detect the release of oxidizable molecules from individual vesicles [4]. Additionally, the number of molecules being released during each exocytosis event can be calculated as well as the kinetics of fusion pore opening and closing. In a whole-tissue preparation, there are numerous cells undergoing exocytosis simultaneously, limiting the spatial resolution of the technique. However, while the resolution required to detect individual release events is not possible; this approach is still able to determine changes in exocytosis from whole-tissue samples in real time in response to tissue stimulation. In this chapter, we will discuss the materials and methods, including aspects of data analysis, for undertaking amperometric measurements from both whole-tissue and single cells, using our own experience with serotonin release in whole-gut preparations [5, 6] and epinephrine release from single chromaffin cells [7–9] as examples. Most primary cultures involve isolating the tissue or organ of interest, digesting it with an enzyme or enzymes to disrupt connective tissue, filtering out the undigested material, and plating the isolated cells on plastic or glass that has been pre-treated to assist with cell adhesion. We have primarily undertaken single--cell amperometry from primary chromaffin cells secreting adrenaline [10–12] and enterochromaffin (EC), cells which secrete serotonin (5-hydroxytryptophan, 5-HT) [13–15]. Other primary cell types and cell lines can alternatively be studied, so long as they secrete an oxidizable amine such as adrenaline, noradrenaline, dopamine, or serotonin. EC cells are the major site of peripheral 5-HT synthesis and secretion. EC cells are located among the epithelial cells lining the gut lumen and, as such, much of the research on EC cell function has focused on their role in the gastrointestinal (GI) tract. Our current knowledge regarding the regulation of

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5-HT secretion from EC cells has largely been gained from studies on intact gastrointestinal tissue. Several groups have successfully measured 5-HT release from the GI tract, using carbon-fiber amperometry. These include measurements from the colon [5, 6] and ileum [16] of various rodent species.

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Materials Chromaffin Cell Culture: 1. Locke’s buffer for tissue digestion (145 mM NaCl, 5.6 mM KCl, 3.6 mM NaHCO3, 5.6 mM glucose, 5.0 mM HEPES, pH 7.4).

2. Collagenase type A, 3 mg/mL in Locke’s buffer. 3. Sterile Dulbecco’s modified Eagle’s medium (DMEM) supplemented with 10% (v/v) heat inactivated fetal calf serum, 100 units/mL penicillin, and 100 mg/mL streptomycin. 4. Refrigerated centrifuge. Gut Tissue Preparations: 1. Krebs solution: 118 mM NaCl, 4.7 mM KCl, 1.0 mM NaHPO4·2H2O, 25 mM NaHCO3, 1.2 mM MgCl·6H2O, 11 mM D-Glucose, 2.5 mM CaCl2·2H2O for dissection, and experiments.

2. Sylgard-lined Petri dish containing oxygenated Krebs solution and dissection microscope. 3. Fine dissection equipment suitable for rodent dissection. Larger dissection equipment may be needed if utilizing human gastrointestinal tissue. For Recordings: 1. Perfusion system, mechanical.

either

gravity

perfusion

system

or

2. Any standard amplifier, digital-to-analogue converter, and relevant software typically used for patch clamp or voltametric recordings. Signal sampling occurs at 10 kHz, using a 1 kHz low-pass hardware filter. Essentially, any patch clamp setup can be used to measure these currents as long as a suitable potential can be provided. 3. A temperature controller through which the solution flows or a heated stage to maintain the experimental bath near physiological temperatures. 4. Carbon-fiber electrodes, typically, 5–10 μm in diameter. Depending on style, they may need to be backfilled with mercury or some other conductive liquid such as 3 M KCl.

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5. A micromanipulator for electrode mounting and positioning of the electrode in close apposition to the recording cell. 6. Light microscope capable of up to 400 magnification for single-cell visualization and placing of electrode onto or near cell surface. 7. This must be located on a vibration–isolated table and inside a Faraday cage so as to remove external electrical background. Grounding wires should be connected to this cage and all electrical equipment within the cage.

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Methods

3.1 Chromaffin Cell Culture

1. Mice are humanely killed and the adrenal glands removed. It is ideal to start with around 6–8 adrenal glands per culture. 2. The adrenal medulla is dissected out from each gland (see Note 1) in ice-cold Locke’s buffer (145 mM NaCl, 5.6 mM KCl, 3.6 mM NaHCO3, 5.6 mM glucose, 5.0 mM HEPES, pH 7.4) [7, 17]. 3. The medullae are then incubated for 30 min in 5 mL Collagenase type A (3 mg/mL in Locke’s buffer) in a shaking water bath set at 37  C. During this time, the suspension is triturated carefully with a pipette at 15 min, 25 min, and 30 min, respectively (see Note 2). 4. The enzyme solution is diluted in ice-cold Locke’s buffer and the cells pelleted at 1000  g for 10 min at 4  C. 5. The cell pellet is resuspended in Dulbecco’s modified Eagle’s medium (DMEM) supplemented with 10% (v/v) heatinactivated fetal calf serum, 100 units/mL penicillin, and 100 mg/mL streptomycin and filtered through a 40-μm nylon mesh. 6. The filtrate is centrifuged as above to pellet cells, which are resuspended in supplemented medium and plated onto pre-treated 35 mm2 plastic tissue culture dishes or coated coverslips, depending on how you will house your samples during the experiment (see Note 3).

3.2 Gastrointestinal Tissue Preparation

1. Mice of either sex are euthanized humanely by inhalation of anesthetic (Nembutal) followed by cervical dislocation. 2. The entire colon is removed and placed in room temperature Krebs solution (118 mM NaCl, 4.7 mM KCl, 1.0 mM NaHPO4·2H2O, 25 mM NaHCO3, 1.2 mM MgCl·6H2O, 11 mM D-Glucose, 2.5 mM CaCl2·2H2O), constantly bubbled with carbogen gas (95% O2/5% CO2).

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3. When working with an open-sheet conformation, a midline incision is made along the mesenteric border and the entire colon pinned with mucosal side uppermost in a Sylgard-lined Petri dish containing oxygenated Krebs solution. 3.3 Amperometric Recordings

1. A suitable oxidizing voltage (750–800 mV for catecholamines, 350–400 mV for serotonin) is applied to the electrode under voltage clamp conditions (see Note 5). 2. Recordings occur by providing this constant potential and recording current changes over a required period of time. 3. Electrodes are mounted on a micromanipulator and placed in close apposition to the recording cell (see Note 4). 4. For recordings of serotonin release from whole-gut tissue in an open-sheet conformation, the whole colon is placed mucosa uppermost in a Sylgard-lined organ bath and continuously perfused with oxygenated buffer at 35–37  C. A voltage of 350–400 mV is passed through the carbon electrode, which is placed just above the tissue surface to avoid contact with the mucosa during colonic contractions (see Note 5). 5. Exocytosis is stimulated in any number of ways, including electrical field stimulation or pharmacologically. This will depend on the cell type being examined, but, in general, all cells will respond to changing the perfusing solution to one containing a 70 mM K+ (see Note 6) [18] (this solution is identical in composition to the standard bath solution but with 70 mM K+ replacing an equimolar amount of NaCl).

3.4

Analysis

1. Current “spikes” due to oxidation of released adrenaline are recorded under baseline and stimulated conditions. In wholetissue recordings, changes in baseline current are used as measures of altered secretion levels. 2. In our laboratory, the Pulse data files are converted to Axon Binary Files and secretory spikes analyzed for the period of stimulation. 3. In single-cell recordings, each spike represents a single exocytosis event (Fig. 1). The parameters which can be analyzed from a single spike include the amplitude, rise time, half width, and decay time. A spike may display what is called a pre-spike foot (PSF) signal that corresponds to the formation of the fusion pore at the plasma membrane or a stand-alone foot signal (SAF), which represents kiss-and-run exocytosis, where the fusion pore closes before it is stable, resulting in no current spike occurring after the initial foot signal [19] (see Note 7). 4. It is important to use a specific threshold of detection, above which exocytosis is thought to have occurred. This is to

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Fig. 1 Single cell amperometry in mouse chromaffin cells. (a) Example amperometric trace from a single cell stimulated (dashed line) with 70 mM K+ solution. (b) A single full fusion event without a pre-spike foot signal, (c) a full fusion event with a pre-spike foot signal and (d) a kiss and run event represented as a stand-alone foot signal are shown. Scale bars in (a) represent 10 s (horizontal) and 100 pA (vertical) and in (b–d) 2 ms (horizontal) and 10 pA (vertical)

overcome signal-to-noise issue with baseline current detection. For single-spike analysis, we select amperometric spikes for analysis if spike amplitude exceeds 10 pA, or at least 2.5 times the root-mean-squared noise of the baseline. We then are able to select the number of single spikes in a selected time period (see Note 8). 5. For kinetic analysis of spikes and PSF signals, only those events that are not overlapping are included. Only PSF signals longer than 1 ms and above 2.5 times the root-mean-squared noise of the baseline are analyzed as foot signals. Rise time of each spike is calculated from the 50% to 90% rising phase, in order to avoid skewing caused by PSF signals (see Note 9). 6. In analyzing spike kinetics, all spikes that meet our threshold criteria are included in calculating the median values of each spike parameter for each cell. The averages of these median values are then used to compare each parameter between cell populations [20]. This is done to avoid errors associated with pooling large numbers of spikes from cells where there is a large cell to cell variability. 7. Analysis of PSF signal kinetics is performed using pooled data from all recorded cells, as many recordings may contain a low number of foot signals. PSF onset is where the signal exceeds the peak-to-peak noise of a 5-ms segment, while the end of the PSF is the inflection point between the PSF signal and the spike. PSF duration is the intervening interval. The PSF area is taken as the integral from the PSF onset to the time where the spike current falls to 2.5 times the root mean squared (RMS) noise of the baseline. 8. Spike kinetic data is nonparametrically distributed and can be evaluated for statistical significance using the Mann-Whitney U test.

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Fig. 2 Amperometric recordings reveal cyclic release of 5-HT during spontaneous contractions in mouse colon [5]

9. For whole-tissue recordings, it is possible to either assess relative changes in current as a gauge of exocytosis or calibrate the electrode before and after an experiment against known concentrations of the analyte being measured [5] (see Note 10). Gut serotonin release can occur in conjunction with muscle contractile events (Fig. 2) or in response to enterochromaffin cell activators such as nutrients.

4

Notes 1. For chromaffin cell cultures, maximizing the ratio of medullary:cortical cells harvested is a trade-off between a higher proportion of medullary cells and an increased preparation time (that will cause increased cell apoptosis). Practice will improve medullary dissection times and cell culture quality. 2. Healthy cells are essential and preferable over high cell yields, given these are single-cell recordings. The trituration step is an essential one and must be performed to a minimum and with great care to avoid cell damage. We feel it is better to have less tissue breakdown in an attempt to have healthier cells. 3. At least 3–4 days of incubation is required for cells to regain function, following the digestions and isolation protocol. This can be possibly longer, depending on the cell type being used. 4. Ensure that the cells are properly attached to the chamber being used for recordings, and that the perfusion system is not so strong that it moves cells during recordings. Cells must remain in the same place during recordings, and, likewise, the probe itself cannot move or drift. If this occurs it can induce artifacts through pressure-evoked release when the cell and probe make contact, or move the probe too far from the cell and lose the spatial fidelity needed to acquire sharp current spikes for consistent recording. 5. Compression-evoked secretion can be induced in whole-gut preparations, [16] but this must be carefully undertaken so as

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to compress the tissue to the same degree in each experiment. Employ a micromanipulator with a digital z-axis reading to ensure this. 6. A carbon-fiber electrode should typically not be used for more than five recordings, to avoid probe desensitization. In order to remove the effects of variance between different probes, comparisons of spike kinetics should preferentially be made by recording from one cell per experimental group with the same electrode. 7. This analysis is employed in our laboratory for the particular equipment setup we use. Many alternative software and analysis packages can be used to the same ends. Further information on amperometric data analysis can be seen elsewhere [3]. 8. There is a high degree of variability in non-neuronal cell amperometric spikes. It is therefore imperative that large data sets are employed so as to avoid type 1 errors that can occur when small numbers of events can skew analysis of underpowered data sets. 9. However, if you decide to analyze your data, it is essential that you remain absolutely consistent across all recordings. Using strict criteria allows the analyzer to avoid problems that can arise during the assessment of such heterogeneous data sets. 10. In whole-tissue preparations, keeping the electrode just above the tissue ensures that no 5-HT release or signal artifacts are evoked by the electrode compressing the tissue. References 1. Kissinger PT, Hart JB, Adams RN (1973) Voltammetry in brain tissue--a new neurophysiological measurement. Brain Res 55:209–213 2. Marcenac F, Gonon F (1985) Fast in vivo monitoring of dopamine release in the rat brain with differential pulse amperometry. Anal Chem 57:1778–1779 3. Mosharov EV, Sulzer D (2005) Analysis of exocytotic events recorded by amperometry. Nat Methods 2:651–658 4. Wightman RM, Jankowski JA, Kennedy RT, Kawagoe KT, Schroeder TJ, Leszczyszyn DJ, Near JA, Diliberto EJ Jr, Viveros OH (1991) Temporally resolved catecholamine spikes correspond to single vesicle release from individual chromaffin cells. Proc Natl Acad Sci U S A 88:10754–10758 5. Keating DJ, Spencer NJ (2010) Release of 5-hydroxytryptamine from the mucosa is not required for the generation or propagation of colonic migrating motor complexes. Gastroenterology 138:659–670

6. Spencer N, Robinson K, Flack B, Zagorodnyuk P, Keating DJ (2011) Mechanisms underlying distension-evoked peristalsis in Guinea pig distal colon: is there a role for enterochromaffin cells? Am J Physiol Gastrointest Liver Physiol 301:G519–G527 7. Yu Y, Chu P-Y, Bowser DN, Keating DJ, Dubach D, Harper I, Tkalcevic J, Finkelstein DI, Pritchard MA (2008) Mice deficient for the chromosome 21 ortholog Itsn1 exhibit vesicle-trafficking abnormalities. Hum Mol Genet 17:3281–3290 8. Zanin MP, Phillips L, Mackenzie KD, Keating DJ (2011) Aging differentially affects multiple aspects of vesicle fusion kinetics. PLoS One 6: e27820 9. Zanin MP, Mackenzie KD, Peiris H, Pritchard MA, Keating DJ (2013) RCAN1 regulates vesicle recycling and quantal release kinetics via effects on calcineurin activity. J Neurochem 124:290–299 10. Keating DJ, Dubach D, Zanin MP, Yu Y, Martin K, Zhao Y-F, Chen C, Porta S, Arbone´s

Amperometry in Single Cells and Tissue ML, Mittaz L, Pritchard MA (2008) DSCR1/ RCAN1 regulates vesicle exocytosis and fusion pore kinetics: implications for down syndrome and Alzheimer’s disease. Hum Mol Genet 17:1020–1030 11. Wen PJ, Osborne SL, Zanin M, Low PC, Wang H-TA, Schoenwaelder SM, Jackson SP, Wedlich-So¨ldner R, Vanhaesebroeck B, Keating DJ, Meunier FA (2011) Phosphatidylinositol(4,5)bisphosphate coordinates actinmediated mobilization and translocation of secretory vesicles to the plasma membrane of chromaffin cells. Nat Commun 2:491 12. Keating DJ, Winter MA, Hemsley KM, Mackenzie KD, Teo EH, Hopwood JJ, Brooks DA, Parkinson-Lawrence EJ (2012) Exocytosis is impaired in mucopolysaccharidosis IIIA mouse chromaffin cells. Neuroscience 227:110–118 13. Raghupathi R, Duffield MD, Zelkas L, Meedeniya A, Brookes SJH, Sia TC, Wattchow DA, Spencer NJ, Keating DJ (2013) Identification of unique release kinetics of serotonin from Guinea-pig and human enterochromaffin cells. J Physiol 591:5959–5975 14. Zelkas L, Raghupathi R, Lumsden AL, Martin AM, Sun E, Spencer NJ, Young RL, Keating DJ (2015) Serotonin-secreting enteroendocrine cells respond via diverse mechanisms to acute and chronic changes in glucose availability. Nutr Metab (Lond) 12:55

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15. Raghupathi R, Jessup CF, Lumsden AL, Keating DJ (2016) Fusion pore size limits 5-HT release from single enterochromaffin cell vesicles. J Cell Physiol 231(7):1593–1600 16. Bertrand PP (2004) Real-time detection of serotonin release from enterochromaffin cells of the Guinea-pig ileum. Neurogastroenterol Motil 16:511–514 17. Maritzen T, Keating DJ, Neagoe I, Zdebik AA, Jentsch TJ (2008) Role of the vesicular chloride transporter ClC-3 in neuroendocrine tissue. J Neurosci 28:10587–10598 18. Kumar R, Corbett MA, Smith NJC, Jolly LA, Tan C, Keating DJ, Duffield MD, Utsumi T, Moriya K, Smith KR, Hoischen A, Abbott K, Harbord MG, Compton AG, Woenig JA, Arts P, Kwint M, Wieskamp N, Gijsen S, Veltman JA, Bahlo M, Gleeson JG, Haan E, Gecz J (2015) Homozygous mutation of STXBP5L explains an autosomal recessive infantile-onset neurodegenerative disorder. Hum Mol Genet 24(7):2000–2010 19. Jackson J, Papadopulos A, Meunier FA, McCluskey A, Robinson PJ, Keating DJ (2015) Small molecules demonstrate the role of dynamin as a bi-directional regulator of the exocytosis fusion pore and vesicle release. Mol Psychiatry 20:810–819 20. Colliver TL, Hess EJ, Ewing AG (2001) Amperometric analysis of exocytosis at chromaffin cells from genetically distinct mice. J Neurosci Methods 105:95–103

Chapter 16 Measurements of Exocytosis by Capacitance Recordings and Calcium Uncaging in Mouse Adrenal Chromaffin Cells Se´bastien Houy, Joana S. Martins, Ralf Mohrmann, and Jakob Balslev Sørensen Abstract Fusion of vesicles with the plasma membrane and liberation of their contents is a multistep process involving several proteins. Correctly assigning the role of specific proteins and reactions in this cascade requires a measurement method with high temporal resolution. Patch-clamp recordings of cell membrane capacitance in combination with calcium measurements, calcium uncaging, and carbon-fiber amperometry allow for the accurate determination of vesicle pool sizes, their fusion kinetics, and their secreted oxidizable content. Here, we will describe this method in a model system for neurosecretion, the adrenal chromaffin cells, which secrete adrenaline. Key words Neurosecretion, Exocytosis, Vesicles, Chromaffin cells, Capacitance measurement, Carbon-fiber amperometry, Patch-clamp, Electrophysiology, Calcium measurements

1

Introduction The release of neurotransmitters or hormones from neurons and neuroendocrine cells depends on rapid exocytosis of vesicles triggered by a sharp increase in intracellular calcium concentration. Adrenal chromaffin cells have served as a crucial model system for understanding the molecular mechanism underlying calciumregulated exocytosis [1–8]. Indeed, many of the molecular players involved in the formation, translocation, docking, priming, and fusion of vesicles have emerged from studies in chromaffin cells [9–20], and detailed studies have helped elucidate the mechanisms of how these molecules act [21–26]. During exocytosis, vesicles fuse with the plasma membrane, releasing their cargo, and vesicular membrane becomes incorporated in the plasma membrane. Vesicle fusion will therefore lead to an increase in cell surface area, which can be monitored electrically as changes in membrane capacitance [27–32].

Florence Niedergang, Nicolas Vitale and Ste´phane Gasman (eds.), Exocytosis and Endocytosis: Methods and Protocols, Methods in Molecular Biology, vol. 2233, https://doi.org/10.1007/978-1-0716-1044-2_16, © Springer Science+Business Media, LLC, part of Springer Nature 2021

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Single-cell membrane capacitance is measured experimentally via the conventional whole-cell patch-clamp technique. This technique relies on a cell-attached configuration where a glass pipette forms a tight seal with giga-ohm resistance, the “gigaseal,” with the cell membrane. Upon further suction, the membrane section under the pipette tip is ruptured, providing a direct and mechanically stable connection between the pipette solution and the cell cytoplasm—the whole-cell configuration. A key advantage of this technique is that it allows for manipulation of the cell’s content while controlling the transmembrane voltage using the voltage-clamp technique. Whole-cell dialysis permits free diffusion of drugs, chelators, and caged compounds into the cytoplasm [27, 33, 34]. The latter has been particularly valuable when studying exocytosis. Optical uncaging of Ca2+ by photolysis of nitrophenyl-EGTA allows to rapidly increase intracellular calcium concentration [35, 36]. Ratiometric calcium measurements can be performed simultaneously by providing Ca2+ indicator fluorescent dyes in the pipette solution [37, 38]. Together, calcium measurements and calcium uncaging provide control over the key variable that triggers exocytosis and regulates upstream priming steps. Membrane capacitance measurements can further be combined with carbon-fiber amperometry, a technique that permits the detection of single-vesicle exocytotic events and the quantification of vesicle content [39]. Moreover, the outstanding temporal resolution allows for real-time kinetics of transmitter release as limited by the fusion pore [40]. In this electrochemical method, a carbonfiber held at a positive constant potential is placed against the surface of a chromaffin cell. This potential is set above the oxidation potential of the molecule of interest—in chromaffin cells adrenaline and noradrenaline. Electric current caused by the transfer of electrons after transmitter oxidation will be observed as amperometric spikes [27, 30, 41–44]. This chapter describes a method to study Ca2+-regulated exocytosis by a combination of capacitance measurements, amperometry, and Ca2+ measurement and uncaging in mouse adrenal chromaffin cells. These techniques allow to detect and quantify secretory events and provide a framework for understanding exocytosis and its underlying molecular machinery.

2

Materials

2.1

Animals

2.2

Cell Culture

Embryonic (E18) or newborn (P0–P2) C57BL6 mice. 1. Culture medium: dissolve 0.55 g of NaHCO3 in 250 ml Dulbecco’s Modified Eagle Medium (DMEM). Filter and add 1 ml of Penicillin (10,000 U/ml)/Streptomycin (10,000 μg/ ml) and 1 ml of insulin–transferrin–selenium–ethanolamine

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(ITSX) supplement. Store at 4
C for up to 3 months. Prior to dissection, warm the necessary volume of medium in an 8% CO2, incubator at 37
C for at least 30 min, with the lid open. 2. Enzyme solution: dissolve 30 mg of L-cysteine in 150 ml DMEM and add 1.5 ml of 100 mM CaCl2 and 1.5 ml of 50 mM EDTA. Filter, make aliquots of 3 ml, and freeze. Store at 20
C for up to 6 months. Before starting the dissection of the adrenal glands, add 20–25 U/ml of papain (Worthington) to the aliquot and place it in an 8% CO2, incubator at 37
C with the lid open. 3. Inactivating solution: dissolve 375 mg of Albumin and 375 mg of trypsin inhibitor in 135 ml of DMEM. Add 15 ml of fetal calf serum, filter, make 3 ml aliquots and freeze. Store at 20
C for up to 6 months. Prior to the dissection of the adrenal glands, place the aliquot in an 8% CO2, incubator at 37
C for at least 30 min with the lid open. 4. 10 Locke’s solution by dissolving 22.5 g of NaCl, 1.04 g of KCl, 0.29 g of NaH2PO4, 0.76 g of Na2HPO4, and 4.5 g of Dglucose in 250 ml milliQ H2O. Adjust pH to 7.0 with NaOH. For cell culture, dilute this solution 10. 5. Extracellular medium: dissolve 2.12 g of NaCl, 0.5 g of glucose, and 0.6 g of HEPES in 200 milliQ H2O. Add 700 μl of 1 M KCl, 500 μl of 1 M CaCl2, and 250 μl of 1 M MgCl2. Adjust pH to 7.2. Add milliQ H2O until a volume of 250 ml is reached (osmolarity should be ~305 mOsm). 6. Intracellular solution: mix 36.2 μl of 2-BP (2-BP: 250 mM Cs-glutamate and 80 mM Cs-Hepes adjusted to pH 7.2 with CsOH) with 8 μl ATP–GTP mix (20 mM Mg-ATP, 3 mM Na-GTP, and 10 mM HEPES at pH 7.2), 2.8 μl of calcium chloride (100 mM) (this value is to reach a high basal calcium concentration solution ~500–600 nM), 5 μl of 80 mM 4K-Nitrophenyl-EGTA, 6.4 μl of dye mix (dye-mix: 5 mM Fura-4F and 5 mM of Mag-Fura-2; note that Fura dyes should be the cell-impermeant potassium or sodium salts), 19.6 μl of filtered water, and 2 μl of vitamin C (40 mM), pH 7.2 (osmolarity adjusted to ~295 mOsm) (see Notes 1–3). 2.3 Equipment/ Instruments

1. Petri dish (35 mm). 2. 6-well cell culture plates. 3. Coverslips (25 mm). 4. pH meter. 5. Osmometer. 6. Borosilicate glass 0.86  1.50  80 mm.

capillaries

with

filament,

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Controller box

Light source

Diode/PMT Camera

Aperture (adjustable) Sample

Monochromator

Apertures (adjustable)

Flash lamp

395nm short-pass filter

Beamsplitter, e.g. 80% transmission/ 20% reflection

Fluorescence filter cube

Fig. 1 The microscope used for measurements. The monochromator and flash-lamps are attached to a dualport epifluorescence condenser; their illumination areas are defined and controlled by their associated apertures. These should be set before performing any experiment (and calibrations) and must not be modified afterward. Ultraviolet (UV) light from the flash-lamp, which is filtered by a 395 nm short-pass filter, and the monochromator light, is combined by an optical beamsplitter (to get enough uncaging light, often 80% of flash light and 20% of monochromator light is used), and is directed toward the filtercube. The filtercube contains a dichroic mirror, which reflects UV and near-UV light from the flash lamp and the monochromator for uncaging and Ca2+-measurements. If cells expressing a fluorophore (EGFP) are used, it is an advantage to use a dichroic mirror with a higher cutoff (e.g., 495 nm), but with efficient reflection ranging down to 340 nm, so that EGFP and Ca2+-imaging can be carried out with the same filter cube. The emission filter is in this case a long-pass filter adapted to the spectrum of EGFP. The area of fluorescent signal detection is controlled by an aperture and the detection area is visualized by a camera. It is convenient to place a red-filter in the transmission light path of the microscope, so that only long wavelength red light is used to visualize the sample. This can be matched with a short-pass filter in front of the photodiode, in order to allow for visualization of the sample while not interfering with Ca2+ measurements

7. Patch pipette puller. 8. Patch pipette coater and polisher. 9. Inverted microscope with a high-numerical apperture oil immersion objective (e.g., 40) with high transmission in the UV range. 10. 2-port epifluorescence condenser with optical beamsplitter allowing for coupling of for example 80% of the flash light and 20% of the monochromator light into the microscope, and with adjustable apertures for both light sources (Fig. 1).

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11. Flash lamp able to deliver a very brief (1–2 ms) pulse of ultraviolet light when triggered by an external voltage command. 12. Monochromator or other light source with external voltagecontrol of wavelength. 13. Perfusion system that can keep the solution level in the bath constant during active perfusion. 14. Patch-clamp amplifier with software lock-in extension and photometry extension. 15. Photodiode/photomultiplier detection system, with adjustable aperture to allow for selection of the detection window. 16. Recording computer. 17. 5 or 10 μm diameter carbon-fiber electrodes insulated and polished [43, 44]. The microscope with attached light sources and detector is shown in the overview in Fig. 1. 2.4

Software

1. Data acquisition software with software lock-in extension and photometry extension. 2. Data analysis software.

3

Methods

3.1 Mouse Chromaffin Cell Culture (Fig. 2)

Here we describe the steps for chromaffin cell primary culture from embryonic (E18) or newborn mice [10]. Capacitance measurements combined with carbon fiber amperometry and intracellular calcium measurement as well as calcium uncaging are performed on these cultured cells. 1. Euthanize the embryonic or newborn mice by decapitation. 2. Using sterilized forceps, peel the skin away from the spine and carefully remove the sheet of connective tissue beneath the skin, without perforating the body cavity. You will be able to clearly see the spine and ribs at this point. 3. Cut the ribs and make a horizontal cross-sectional cut of the spine. With one set of forceps hold the animal’s body in place and with the other pair grab the spine and pull it toward you. Be sure to clear the spine from any connective tissue before peeling back the spine. 4. Visualize the kidneys and identify the adrenal glands above them. Harvest the glands and place them quickly in a 35 mm petri dish with cold Locke’s solution (see Note 4). Remove any excess tissue with fine sterilized scissors/forceps.

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Fig. 2 Adrenal glands extraction. (a) Peel the skin from a P0–2 mouse and remove any connective tissue. (b, c) Insert the forceps between the ribs and cut the bones by applying pressure on the forceps. (d) Proceed to a horizontal cross-sectional cut of the spinal cord. (e, f) Pull the spinal cord toward the tail and remove it. The adrenal glands will be located above the kidneys (black arrows). (g) Harvest the glands with forceps and place them in a petri dish containing Locke’s solution. (h) Remove any excess tissue from the extracted glands with forceps or fine scissors

5. Aspirate the glands with a 1 ml micropipette and place them in a 1.5 ml Eppendorf tube. Rapidly remove the Locke’s solution and add 200 μl of prewarmed Enzyme solution. Incubate for 30–45 min at 37
C, 8% CO2 (see Note 5). Add 200 μl of prewarmed Inactivating solution and incubate another 5–15 min at room temperature. 6. Meanwhile prepare 6-well plates according to the number of animals euthanized. Place three to four 25 mm coverslips for each animal (see Note 6). 7. After incubation, remove most of the medium with a 1 ml micropipette and wash it once with 200 μl of prewarmed cell culture medium. 8. Exchange the 200 μl of cell culture medium for 180 μl of prewarmed cell culture medium and gently triturate the glands with a 200 μl micropipette by aspirating up and down. Once the gland’s capsule bursts, do a few more slow strokes to release and dissociate the chromaffin cells from the medulla. 9. Plate a 60 μl drop of cell suspension at the center of the coverslip and incubate for 30–45 min at 37
C, 8% CO2. The point of this step is to make the cell attach in a concentrated area in the middle of the coverslip.

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10. Gently add 2 ml of prewarmed cell culture medium to each of the wells and place the plate(s) back in the incubator for 2–3 days. 3.2 How to Set Up for Measurements

1. After inserting a new bulb in the monochromator, follow the instruction of the vendor to calibrate the bulb position and ensure maximum light output. This might involve using a photodiode arrangement connected to a voltmeter and adjustment of orientation screws until the maximum light output is achieved. The same is done after inserting a new bulb into the flash lamp (some instruments do not require this step). 2. To control the wavelength of the light emitted by the monochromator with your preferred software, a calibration curve (straight line) linking the input selection voltage to the output light wavelength must be determined. In some cases, the calibration is provided by the vendor; otherwise, you might use two narrow band-pass filters (we have used 305 and 505 nm filters) to search for the voltage giving maximal light output at each of these two wavelengths. Linear regression is then used to determine the calibration curve. 3. The adjustment of the area of monochromator illumination has to be carried out by adjusting the aperture placed on the epifluorescence condenser (see also Fig. 3). For this purpose, place a glass coverslip marked on the top side with a fluorescent highlighter in the electrophysiological chamber and on the microscope stage, using the immersion objective you are going to use for measurement. Turn on the monochromator and adjust the aperture to determine the monochromator illumination so that it is slightly larger than the largest cell you are going to measure from. 4. The next step is to adjust the area of the excitation light emitted by the flash lamp, which allows for UV-flash uncaging of compounds (Fig. 3). On the flashlamp, use the focus mode, which will cause the flashlamp to flash repetitively at low intensity. Alternatively, place a small lamp with constant light output in the illumination path. Observe the illumination area on the “magic marker” coverslip (use a camera to detect the illumination area—do not look into the microscope while flashing the flash lamp). Adjust the area of the flash lamp illumination with its associated aperture. Make this area slightly larger than the illumination area of the monochromator, but limit the overlap with the carbon fiber to minimize the appearance of an electrical artifact when you flash the lamp. 5. Next, adjust the Photodiode detection area using the aperture (Fig. 1). The area should be large enough to ensure that the entire chromaffin cell is inside the detection area.

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Fig. 3 Chromaffin cell with patch pipette and amperometry electrode. Example of the experimental configuration during a whole-cell patch-clamp recording of a chromaffin cell, 2 days after cell culture. To the left, the 5 μm diameter carbonfiber used to perform the amperometric recording, held at +700 mV; to the right, the patch pipette containing the intracellular solution, which includes calcium cages and calcium dyes, used to record cell membrane capacitance. The dashed rectangles represent the Flash lamp excitation area (FL), in red, the Monochromator excitation area (Mn), in blue, and the photodiode detector area (PD), in green (refer to Fig. 1)

6. For Fura measurements of intracellular Ca2+ concentrations, a Fura dye suitable for the measured concentration range has to be selected. Alternatively, a combination of two Fura dyes can be used, in order to cover a larger concentration range [37, 38]. Using Fura-4F (Kd ¼ 1 μM) and furaptra (Kd ¼ 40 μM) allows for an uninterrupted measurement range from ~100 nM to ~100 μM, which includes both the sub-μM range relevant for vesicle priming [21, 37], and the supra-μM range, which is relevant for fusion triggering. When using a mixture of two dyes, it is essential that the ratio between the concentrations is invariant; any pipetting error will compromise measurements. Therefore, the two dyes have to be premixed and aliquoted (e.g., at a 5/5 mM concentration) before calibration is carried out using the dye mixture (step 7). Measurements of Ca2+ concentrations can be carried out by oscillating excitation wavelengths between 340 nm (350 nm might yield a better signal-to-noise ratio for objectives with limited transmission in the UV range) and 380 nm. 7. A calcium calibration in the cell has to be performed every time a new mix of calcium dyes is taken into use, or after exchange of the bulb of the monochromator (see Note 7). To this end, prepare several (we use step 8) different intracellular solutions containing known calcium concentrations (In our lab, the solutions are prepared with the following calcium concentrations: 0; 331 nM; 741 nM; 1.77 μM; 11.86 μM; 23.74 μM; 72.77 μM; and 10 mM), the calcium dyes and the calcium buffers 1,2,-bis

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(2-aminophenoxy)ethane-N,N-N0 ,N0 -tetraacetate (BAPTA) or 1,3-diaminopropane-2-ol-N,N0 -tetraacetate (DPTA). The total concentration of buffer should be at least 20 mM and the Kd for BAPTA is assumed to be 221 nM, and the Kd of DPTA 81 μM [37]. It is advisable to prepare stock solutions of BAPTA, Ca-BAPTA, DPTA, and Ca-DPTA (e.g., each at 100 mM), which include a little HEPES, such that their pH can be adjusted to 7.2 before aliquoting. The solutions are then prepared by combining BAPTA with Ca-BAPTA (for calibration solutions with Ca2+ concentration below 1 μM) or DPTA with Ca-DPTA (for solutions with Ca2+ above 1 μM). This avoids adding Ca2+ to unbound Ca2+ buffers, which can result in large pH-changes. The calibration solutions should also contain any nucleotides or Mg2+ used during measurements. For each solution, patch 3–5 cells in the whole-cell configuration and allow 5–10 min. For infusion of the solution while ideally maintaining an access resistance 1 μM, the cell might succeed in maintaining a lower Ca2+ concentration for a while, until it is overwhelmed and the ratio increases to a stable value. Use the average of the ratio measured from 3–5 cells at each Ca2+ concentration to construct a calibration curve. 3.3 Whole-Cell Patch-Clamp Capacitance Measurement Combined with Carbon Fiber Amperometry and Intracellular Calcium Concentration Measurement and Calcium Uncaging

1. Pull the patch-clamp pipettes from borosilicate glass capillaries with filament and polish the tip to a final resistance of approximately 3–5 MΩ. Coat the pipette with Sylgard, a hydrophobic silicone [45]. Following coating the Sylgard needs to be cured, for instance by a hot stream of air, and/or by baking in an oven. Coating the pipette has several advantages: it reduces the capacitance of the patch pipette and helps to have an accurate fast capacitance transient compensation; it reduces noise and increases the accuracy of the capacitance measurement [32]; and it protects against capacitance “creeping” due to the bath solution moving up on the outside of the pipette, which might increase the pipette capacitance over time. Polishing the pipette might help to obtain a better seal. 2. Capacitance measurements are conveniently carried out employing a lock-in amplifier implemented in software (see Note 8). Ensure that the software implements the Lindau– Neher method [46]. The parameters of the sinewave have to be chosen before the start of measurements. This includes the frequency and amplitude of the sinewave-shaped voltage command, the number of points per sine-wave cycle, and the reversal potential of the leak current. Refer to the literature for tips [32, 47]. For chromaffin cells, we use a sine-wave frequency around 1000 Hz, and a peak amplitude of 35 mV around a

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Fig. 4 Electrophysiological chamber. Cultured chromaffin cells plated on the coverslip are covered in extracellular medium. The perfusion of the extracellular medium and the aspiration are positioned on opposite sides of each other, to ensure a continuous perfusion of the entire coverslip. The fluid level system detector is placed in contact with the surface of the extracellular medium and is connected to the pump that drives aspiration, such that a constant bath level is maintained. The carbon-fiber electrode and the patch-clamp pipette are positioned in direct contact with the cell with the help of micromanipulators

holding potential of 70 mV, with 12 point per cycle (input and output). The reversal potential is set to 0 mV (see Note 9). 3. Place the coverslip in the electrophysiological chamber and, with a glass Pasteur pipette, quickly add extracellular medium to prevent the cells from drying. Place fluid level controller, reference electrode, perfusion system and aspiration capillaries in contact with extracellular medium in the electrophysiological chamber (Fig. 4, see Note 10). 4. Place a cell in the photodiode detection field (Fig. 3). Use a pipette with a long loading tip to load a small amount of intracellular solution (0.1–0.5 μl) into the tip of the pipette. Mount the pipette on the pipette holder and place it in the bath while maintained at a positive pressure and at a clamp voltage of zero. Adjust the voltage offset of the patch-clamp amplifier so that the amplifier reads zero current (refer to a patch-clamp manual or an introductory text for a complete description of the technique, for instance [45]). Apply a repetitive 5-mV amplitude square pulse, to allow for inspection of resistance and capacitative currents. Using micromanipulators, place the patch pipette so that it touches the cell and remove the positive pressure in the pipette, which usually starts the seal formation. Use gentle suction to achieve the cell-attached configuration and establish a gigaohm seal (see Notes 11 and 12). Gradually decreasing the holding potential in the pipette toward 70 mV will often facilitate giga-seal formation. When this is achieved,

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Fig. 5 High time-resolution measurement chromaffin cells. Top panel: [Ca2+]i before and after UV flash photolysis of the calcium cages (at arrow, see time axis). [Ca2+]i increases from the submicromolar range to ~20 μM after stimulation. Second panel: Left: Capacitance trace over time. Kinetic analysis identifies fast and slow components (inserted to the right). Third panel, left ordinate: recorded amperometric current. A peak in the oxidation current is observed after calcium uncaging. Right ordinate: The integration of the recorded amperometry trace provides the charge of the released content. If only catecholamine-containing vesicles fuse, the integrated amperometry is expected to mimic the capacitance change, except for a diffusional delay. Fourth panel: estimated membrane conductance (Gm). Fifth panel: estimated series/access conductance (Gs)

adjust the pipette holding potential to 70 mV (not corrected for liquid junction potential). Cancel the fast capacitive transient currents caused by stray capacitance of the pipette and pipette holder using the “fast” capacitative cancellation circuit of the amplifier. 5. At this stage, while still in the cell-attached configuration, wait until the bath perfusion has removed any traces of fura dyes, which have leaked out of the pipette during seal formation.

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Then, perform a control fluorescence measurement at wavelength 340 nm (or 350 nm) and 380 nm. In practice, apply repetitive fluorescence measurements in the cell-attached situation, until the readings stabilize, indicating that the Fura dyes have been washed away. The control fluorescence measurement accounts for the tip of the dye-filled pipette, which is placed inside the detection area. The values from the control fluorescence measurement should be subtracted from fluorescence values later obtained in whole-cell configuration. It is important that the pipette and cell do not move with respect to the detection window after the control fluorescence measurement has been carried out (see Note 13). 6. With strong, but brief, suction, rupture the patch of membrane to achieve the whole-cell configuration. The appearance of slow current transients marks the establishment of the wholecell configuration. Cancel the capacitance transients using the slow capacitance cancellation circuit. If using an automatic fitting-routine, repeat a few times until the cancellation is optimal. Note that access resistance compensation is not used together with capacitance measurements [32]. 7. Keep the whole-cell configuration for at least 30 s to allow the calcium dyes and the calcium cage time to diffuse into the cell (see Note 14). Use this time period to place the carbon-fiber electrode in contact with the cell, using a second micromanipulator. Carbon-fiber electrodes are held at a potential of 700–800 mV using a separate patch-clamp amplifier, currents are low-pass filtered with a 7-pole Bessel filter at 1 or 3 kHz and sampled at 11.5 kHz. 8. Trigger cell secretion by triggering UV-flash photolysis of the calcium cage. An example of a Ca2+ uncaging measurement is shown in Fig. 5 (see Notes 15–17). Alternatively, stimulate using a depolarization protocol; in this case nitrophenylEGTA should obviously not be included in the intracellular solution, and the total Ca2+ buffer capacity should be set using the Ca2+ dyes and EGTA (0.5–1 mM). Several useful depolarization and UV-flash photolysis protocols can be found in the literature, which help distinguish different vesicle pools, as for instance the Immediately Releasable Pool (IRP; [48, 49]), the Readily Releasable and the Slowly Releasable Pool (RRP and SRP; [35, 37]), and the High-Calcium Sensitive Pool (HCSP; [50]). Gradual release of Ca2+ from nitrophenyl-EGTA using low-level illumination can create a Ca2+-ramp, which is useful for measuring the Ca2+ threshold for secretion [38]. 9. Save the file as an individual recording for further analysis and create a new file to record a new cell. 10. Repeat steps 1–9. A lot.

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Notes 1. The Kd of nitrophenyl-EGTA for Ca2+ is 80 nM [36]. It is important to ensure that the concentration of free calcium in the intracellular solution is above this value. Otherwise, the unbound nitrophenyl-EGTA will act as an extra buffer, which will counteract Ca2+-increases. Other Ca2+ cages are available, with variable photochemical yield, effectiveness of light absorption, rate of Ca2+ release, pH sensitivity, and affinities and selectivities for Ca2+ over Mg2+ [51]. 2. Because nitrophenyl-EGTA is an EGTA-derivative, Ca2+ binding is pH-dependent [51]. This has practical consequences: when adding Ca2+ to a pH-neutral solution of nitrophenylEGTA, the pH will drop precipitously, due to release of protons. To avoid this, dissolve the nitrophenyl-EGTA tetrapotassium salt in water, but do not pH-adjust the solution, which will be alkaline. When assembling the pipette solution, Ca2+ addition to the alkaline nitrophenyl-EGTA solution will help neutralize the pH. In addition, the pipette solution should contain large amounts (30–40 mM) of a fast pH-buffer such as 2-[4-(2-hydroxyethyl)piperazin-1-yl]ethanesulfonic acid (HEPES) adjusted to pH ~7.2. This will counteract pH changes when nitrophenyl-EGTA is broken down by the flash light. Check the pH of the assembled pipette solution by spotting a small drop on a piece of pH-paper. 3. When assembling the intracellular solution, unavoidable pipetting errors due to the very small volumes used (we usually prepare 40 or 80 μl solution at a time) can lead to a solution with a [Ca2+] slightly off target. Have a solution of nitrophenyl-EGTA (10–20 mM) and another of CaCl2 (10–20 mM) ready at the setup, so you can add a little of either one (0.1–0.2 μl at a time) to the intracellular solution, in order to correct the [Ca2+]. 4. When performing primary cell culture, do not leave the glands in Locke’s solution for a long period of time. Be as quick as possible. During trituration of the adrenal glands, perform slow strokes without creating any bubbles. Once the gland capsules burst, do not overtriturate. Perform one or two additional strokes. 5. Before cell culture, it is important that the enzyme solution is equilibrated for at least 1 h at 8% CO2. 6. Note that chromaffin cells like company: even if only a few cells can be patched on each coverslip, the cell culture should be relative dense to keep the cells healthy. Therefore, chromaffin cells isolated from a single mouse should not be spread across more than four coverslips. The viability of the cells influences

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sealing and maintenance of the patch-clamp configuration. Usually, experiments should be performed 2–4 days after cell culture. 7. Calcium calibration should be accurately performed before starting any experiment to validate that you get an accurate pre- and postflash calcium concentration. A wrong calibration can lead to critical variation in the burst size, kinetics of vesicle release, or even the absence of secretion. 8. It is often useful to set up the detection software to save values from intermittent calcium and capacitance measurements throughout the experiment, such that in the end overview of the entire experiment can be produced (Fig. 6), including the fluorescence signals at 340 nm (or 350 nm) and 380 nm, the fluorescence ratio, the Ca2+ concentration, the access conductance, leak current and membrane capacitance. This gives a convenient overview of the experiment, to confirm that parameters (leak, access and Ca2+ concentrations) are within the desired ranges. It also helps to establish whether capacitance was changing before uncaging or depolarization, indicating the fusion of vesicles throughout the experiment, which might otherwise go undetected. 9. The Cm output of the lock-in amplifier is usually a good estimate of capacitance, provided the cell is small and round (so that it can be modeled by a single compartment RC-circuit, [32]) and the membrane resistance, Rm, is very high compared to the access resistance, Ra. Overall, single chromaffin cells are near-optimal specimens for capacitance measurements. Nevertheless, it is advisable to inspect the estimates of Ra for changes that are correlated with Cm-changes, as these can be indicative of an incorrect phase setting, or a cell not well modeled by a single compartment circuit [32]. Refer to [32] for a full treatment of the capacitance measurement techniques and its limitations and pitfalls. 10. For optimal recording conditions, frequently oxidize the silver wire of your recording electrode by submerging it in chlorine solution or electroplating it with hydrochloric acid. Use a Ag/ AgCl-pellet as reference electrode. If combining capacitance and amperometry measurements, use a single reference electrode attached to the “ground” input of both preamplifiers. 11. If the cell-attached configuration does not form easily: (a) Check the osmolarity of pipette and bath solution. For optimal seal formation, the extracellular solution should be 5–10 mOsm hyperosmolar compared to the intracellular solution. (b) Absence of positive pipette pressure can prevent you from achieving the gigaseal and can lead to aspiration of bath fluid into the pipette. To confirm the positive pressure (around

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Fig. 6 Low time-resolution overview of the experiment. Overview of the different patch-clamp parameters recorded throughout an experiment, where the cell was stimulated three times (marked 1–3) by calcium uncaging. The traces represented are, from top to bottom, the fluorescence signals of the Fura dyes at 380 nm (black) and 350 nm (green), the ratio of the fluorescence signal 350 nm/380 nm, the calcium concentration ([Ca2+]i), the series/access conductance (Gs), the leak current (pA), and the cell membrane capacitance (Cm). Note that there are two Gs and Gm-traces. This is because the slow capacitance cancellation circuit gives one value, which is only updated when the capacitance cancellation is repeated, whereas the Lindau–Neher technique gives another value, which is updated more frequently throughout the experiment. In order to generate the low time-resolution overview, we execute a small command, consisting of a 100-ms sinewave together with a 350 and 380 nm fluorescence measurement at low frequency (0.2–0.5 Hz) throughout the experiment

10 cm H2O), add dye to the patch pipette and put it in the bath. In the presence of positive pressure, the dye will flow into the bath. If no positive pressure is found, check the tubing for holes, and replace the O-ring holding the pipette in the pipette holder. (c) Check the cell quality. This is based on experience, but chromaffin cells should be round and have good contrast in the microscope. Loss of contrast or a distorted shape is indicative of a problem. A low cell density is also problematic (see above). 12. During patch formation suction should be gentle and limited so that the pipette compounds are not aspirated and not too much membrane is sucked into the pipette, which can make it hard to obtain the whole-cell configuration later. It is often possible to form the cell-attached configuration while applying almost no suction.

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13. Instead of a photodiode/photomultiplier (Fig. 1) a camera could be used to detect the fluorescence. In this case, the camera software should be set up such that the 340 nm/ 380 nm fluorescence-ratio (corrected for values obtained in the cell-attached configuration, see Subheading 3.3, step 5) is calculated and displayed online during the experiment. This will allow for the determination of the prestimulus [Ca2+]i before the uncaging flash is given. 14. The prestimulus [Ca2+]i controls vesicle priming and thereby the size of the vesicle pools in chromaffin cells [37]; it is therefore an essential parameter to measure and control when doing pool size measurements. To do so, use an intracellular solution with a slightly lower [Ca2+] than desired. Upon infusion, increase the resting [Ca2+]i by low intensity and fast oscillating monochromator illuminations at 340 nm (or 350 nm) and 380 nm, which will induce slow Ca2+ uncaging while at the same time allowing you to measure [Ca2 + ]. Monitor the 340/380 ratio online and compare with your calibration, until you reach the desired prestimulus [Ca2+]i. Reduce the monochromator illumination such that the [Ca2 + ]i is maintained for at least 10–20 s at that level before applying a flash [37]. 15. If no or too-low secretion is unexpectedly observed, ensure that the patched cell is entirely inside the UV flash lamp illumination field, that the resting Ca2+ concentration is neither too low (this can lead to suboptimal vesicle priming, see below) nor too high (this could lead to premature fusion of vesicles before the uncaging measurements can be carried out; keep an eye on the amperometric and capacitance recording while loading the cell, as this can reveal whether vesicles are fusing), that the flash illumination is sufficiently strong to photolyze the cage (i.e., measure [Ca2+] after a flash), that the pH of the pipette solution is approximately neutral, that the chromaffin cells look healthy, and that the bath solution is correct. 16. If the carbon fiber is unexpectedly not detecting any events, cutting and polishing the fiber may increase sensitivity. Inversely polarizing the electrode may also help to regain sensitivity. 17. While it is not obligatory to perform capacitance and amperometric measurements in parallel, comparing the capacitance and the amperometric signal allows for the further determination that the vesicles fusing (as detected by capacitance increases) actually released adrenaline/noradrenaline (as detected by the amperometric current), as expected for dense-core vesicles. Under certain circumstances, capacitance increases not related to adrenaline/noradrenaline-release can

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be detected [52]. In comparing capacitance and amperometry it should be remembered that the latter method only detects release from a part of the cell, after a diffusional delay. References 1. Sorensen JB (2004) Formation, stabilisation and fusion of the readily releasable pool of secretory vesicles. Pflugers Archiv 448:347–362 2. Verhage M, Sorensen JB (2008) Vesicle docking in regulated exocytosis. Traffic 9:1414–1424 3. Neher E (2018) Neurosecretion: what can we learn from chromaffin cells. Pflugers Archiv 470:7–11 4. Rettig J, Neher E (2002) Emerging roles of presynaptic proteins in Ca++-triggered exocytosis. Science 298:781–785 5. Marengo FD, Cardenas AM (2018) How does the stimulus define exocytosis in adrenal chromaffin cells? Pflugers Arch 470:155–167 6. Dhara M, Mohrmann R, Bruns D (2018) v-SNARE function in chromaffin cells. Pflugers Arch 470:169–180 7. Stevens DR, Schirra C, Becherer U, Rettig J (2011) Vesicle pools: lessons from adrenal chromaffin cells. Front Synaptic Neurosci 3:2 8. Bader MF, Holz RW, Kumakura K, Vitale N (2002) Exocytosis: the chromaffin cell as a model system. Ann N Y Acad Sci 971:178–183 9. Steyer JA, Horstmann H, Almers W (1997) Transport, docking and exocytosis of single secretory granules in live chromaffin cells. Nature 388:474–478 10. Sorensen JB, Nagy G, Varoqueaux F, Nehring RB, Brose N, Wilson MC, Neher E (2003) Differential control of the releasable vesicle pools by SNAP-25 splice variants and SNAP23. Cell 114:75–86 11. Borisovska M, Zhao Y, Tsytsyura Y, Glyvuk N, Takamori S, Matti U, Rettig J, Sudhof T, Bruns D (2005) v-SNAREs control exocytosis of vesicles from priming to fusion. EMBO J 24:2114–2126 12. Voets T, Moser T, Lund PE, Chow RH, Geppert M, Sudhof TC, Neher E (2001) Intracellular calcium dependence of large densecore vesicle exocytosis in the absence of synaptotagmin I. Proc Natl Acad Sci U S A 98:11680–11685 13. Voets T, Toonen RF, Brian EC, de Wit H, Moser T, Rettig J, Sudhof TC, Neher E, Verhage M (2001) Munc18-1 promotes large

dense-core vesicle docking. Neuron 31:581–591 14. Schonn JS, Maximov A, Lao Y, Sudhof TC, Sorensen JB (2008) Synaptotagmin-1 and -7 are functionally overlapping Ca2+ sensors for exocytosis in adrenal chromaffin cells. Proc Natl Acad Sci U S A 105:3998–4003 15. Liu Y, Schirra C, Stevens DR, Matti U, Speidel D, Hof D, Bruns D, Brose N, Rettig J (2008) CAPS facilitates filling of the rapidly releasable pool of large dense-core vesicles. J Neurosci 28:5594–5601 16. Speidel D, Bruederle CE, Enk C, Voets T, Varoqueaux F, Reim K, Becherer U, Fornai F, Ruggieri S, Holighaus Y, Weihe E, Bruns D, Brose N, Rettig J (2005) CAPS1 regulates catecholamine loading of large dense-core vesicles. Neuron 46:75–88 17. Man KN, Imig C, Walter AM, Pinheiro PS, Stevens DR, Rettig J, Sorensen JB, Cooper BH, Brose N, Wojcik SM (2015) Identification of a Munc13-sensitive step in chromaffin cell large dense-core vesicle exocytosis. eLife 4: e10635 18. Ashery U, Varoqueaux F, Voets T, Betz A, Thakur P, Koch H, Neher E, Brose N, Rettig J (2000) Munc13-1 acts as a priming factor for large dense-core vesicles in bovine chromaffin cells. EMBO J 19:3586–3596 19. Cai H, Reim K, Varoqueaux F, Tapechum S, Hill K, Sorensen JB, Brose N, Chow RH (2008) Complexin II plays a positive role in Ca2+-triggered exocytosis by facilitating vesicle priming. Proc Natl Acad Sci U S A 105:19538–19543 20. Vitale ML, Seward EP, Trifaro JM (1995) Chromaffin cell cortical actin network dynamics control the size of the release-ready vesicle pool and the initial rate of exocytosis. Neuron 14:353–363 21. Houy S, Groffen AJ, Ziomkiewicz I, Verhage M, Pinheiro PS, Sorensen JB (2017) Doc2B acts as a calcium sensor for vesicle priming requiring synaptotagmin-1, Munc132 and SNAREs. eLife 6:e27000 22. Sorensen JB, Wiederhold K, Muller EM, Milosevic I, Nagy G, de Groot BL, Grubmuller H, Fasshauer D (2006) Sequential N- to C-terminal SNARE complex assembly

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drives priming and fusion of secretory vesicles. EMBO J 25:955–966 23. Mohrmann R, de Wit H, Verhage M, Neher E, Sorensen JB (2010) Fast vesicle fusion in living cells requires at least three SNARE complexes. Science 330:502–505 24. Shaaban A, Dhara M, Frisch W, Harb A, Shaib AH, Becherer U, Bruns D, Mohrmann R (2019) The SNAP-25 linker supports fusion intermediates by local lipid interactions. eLife 8:e41720 25. Makke M, Mantero Martinez M, Gaya S, Schwarz Y, Frisch W, Silva-Bermudez L, Jung M, Mohrmann R, Dhara M, Bruns D (2018) A mechanism for exocytotic arrest by the Complexin C-terminus. eLife 7:e38981 26. Mohrmann R, de Wit H, Connell E, Pinheiro PS, Leese C, Bruns D, Davletov B, Verhage M, Sorensen JB (2013) Synaptotagmin interaction with SNAP-25 governs vesicle docking, priming, and fusion triggering. J Neurosci 33:14417–14430 27. Angleson JK, Betz WJ (1997) Monitoring secretion in real time: capacitance, amperometry and fluorescence compared. Trends Neurosci 20:281–287 28. Khvotchev M, Kavalali ET (2008) Pharmacology of neurotransmitter release: measuring exocytosis. Handb Exp Pharmacol 184:23–43 29. Neher E, Marty A (1982) Discrete changes of cell-membrane capacitance observed under conditions of enhanced secretion in bovine adrenal chromaffin cells. Proc Natl Acad Sci U S A 79:6712–6716 30. Borges R, Camacho M, Gillis KD (2008) Measuring secretion in chromaffin cells using electrophysiological and electrochemical methods. Acta Physiol 192:173–184 31. Lindau M, Neher E (1988) Patch-clamp techniques for time-resolved capacitance measurements in single cells. Pflugers Arch 411:137–146 32. Gillis KD (1995) Techniques for membrane capacitance measurements. In: Sakmann B, Neher E (eds) Single-channel recording, 2nd edn. Plenum Press, New York, pp 155–198 33. Segev A, Garcia-Oscos F, Kourrich S (2016) Whole-cell patch-clamp recordings in brain slices. J Vis Exp 112:54024 34. Conforti L (2012). Chapter 20: patch-clamp technique. In: Sperelakis N (ed) Cell physiology source book, 4th edn. Academic, Cambridge 35. Heinemann C, Chow RH, Neher E, Zucker RS (1994) Kinetics of the secretory response in bovine chromaffin cells following flash photolysis of caged Ca2+. Biophys J 67:2546–2557

36. Ellis-Davies GC, Kaplan JH (1994) Nitrophenyl-EGTA, a photolabile chelator that selectively binds Ca2+ with high affinity and releases it rapidly upon photolysis. Proc Natl Acad Sci U S A 91:187–191 37. Voets T (2000) Dissection of three Ca2+dependent steps leading to secretion in chromaffin cells from mouse adrenal slices. Neuron 28:537–545 38. Sorensen JB, Matti U, Wei SH, Nehring RB, Voets T, Ashery U, Binz T, Neher E, Rettig J (2002) The SNARE protein SNAP-25 is linked to fast calcium triggering of exocytosis. Proc Natl Acad Sci U S A 99:1627–1632 39. Wightman RM, Jankowski JA, Kennedy RT, Kawagoe KT, Schroeder TJ, Leszczyszyn DJ, Near JA, Diliberto EJ Jr, Viveros OH (1991) Temporally resolved catecholamine spikes correspond to single vesicle release from individual chromaffin cells. Proc Natl Acad Sci U S A 88:10754–10758 40. Chow RH, von Ruden L, Neher E (1992) Delay in vesicle fusion revealed by electrochemical monitoring of single secretory events in adrenal chromaffin cells. Nature 356:60–63 41. Fathali H, Cans AS (2018) Amperometry methods for monitoring vesicular quantal size and regulation of exocytosis release. Pflugers Archiv 470:125–134 42. Mosharov EV, Sulzer D (2005) Analysis of exocytotic events recorded by amperometry. Nat Methods 2:651–658 43. Bruns D (2004) Detection of transmitter release with carbon fiber electrodes. Methods 33:312–321 44. Chow RH, Ru¨den L (1995) Electrochemical detection of secretion from single cells. In: Sakmann B, Neher E (eds) Single-channel recording, 2nd edn. Plenum Press, New York 45. Penner R (1995) A practical guide to patch clamping. In: Sakmann B, Neher E (eds) Single-channel recording, 2nd edn. Plenum Press, New York 46. Lindau M, Neher E (1988) Patch-clamp techniques for time-resolved capacitance measurements in single cells. Pflugers Archiv 411:137–146 47. Chen P, Gillis KD (2000) The noise of membrane capacitance measurements in the wholecell recording configuration. Biophys J 79:2162–2170 48. Horrigan FT, Bookman RJ (1994) Releasable pools and the kinetics of exocytosis in adrenal chromaffin cells. Neuron 13:1119–1129 49. Voets T, Neher E, Moser T (1999) Mechanisms underlying phasic and sustained secretion

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Chapter 17 Retention Using Selective Hooks-Synchronized Secretion to Measure Local Exocytosis Gaelle Boncompain, Lou Fourriere, Nelly Gareil, and Franck Perez Abstract Proteins destined to be exposed to the extracellular space enter the secretory pathway at the level of the endoplasmic reticulum. Proteins are then transported to the Golgi apparatus and addressed to their destination compartment, such as the plasma membrane for exocytic cargos. Exocytosis constitutes the last step of the anterograde transport of secretory cargos. Exocytic vesicles fuse with the plasma membrane, releasing soluble proteins to the extracellular milieu and transmembrane proteins to the plasma membrane. In order to monitor local exocytosis of cargos, we describe in this chapter how to perform synchronization of the anterograde transport of an exocytic cargo of interest using the retention using selective hooks (RUSH) assay in combination with selective protein immobilization (SPI). SPI is based on the coating of coverslips with anti-green fluorescent protein (GFP) antibodies, which capture the GFP-tagged RUSH cargos once exposed to the cell surface after its release by the addition of biotin. Key words Exocytosis, Secretory transport, RUSH, Antibody coating, Protein immobilization, Realtime imaging

1

Introduction Exocytosis is the last step of the anterograde transport of proteins destined to the cell surface through the secretory pathway. It represents an essential step to ensure cell homeostasis such as the release of growth factors, the deposition of extracellular matrix components, or the expression of surface receptors. The exocytosed cargos can be proteins either soluble, transmembrane, or glycosylphosphatidylinositol (GPI)-anchored. At steady state, exocytic events observed at the cell surface originate either from anterograde transport or from recycling pathways. To analyze events of exocytosis linked to the anterograde transport of the cargo of interest and distinguish them from the recycling pathway, the best method is to synchronize cargo transport along the secretory pathway. Several years ago, we developed the Retention Using Selective Hooks (RUSH) assay to synchronize the anterograde transport of virtually

Florence Niedergang, Nicolas Vitale and Ste´phane Gasman (eds.), Exocytosis and Endocytosis: Methods and Protocols, Methods in Molecular Biology, vol. 2233, https://doi.org/10.1007/978-1-0716-1044-2_17, © Springer Science+Business Media, LLC, part of Springer Nature 2021

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any cargo of interest in physiological conditions for cells [1]. The RUSH assay is a two-state secretory assay based on the retention of the cargo of interest (reporter) in a donor compartment and its release to the acceptor compartment. The donor compartment is usually the endoplasmic reticulum (ER) when using the RUSH assay, while various final compartments can be targeted such as various domains of the plasma membrane, the endosomes, or the lysosomes, for example. Here, we will focus on the study of transport to the cell surface and exocytosis (Fig. 1a). Retention of the reporter is mediated by the interaction between streptavidin fused to an ER-resident hook and a streptavidin-binding peptide (SBP) fused to the reporter. Interaction is then released by addition of biotin. Biotin has a higher affinity for streptavidin than SBP [2, 3] competing out the SBP. The reporter is then free to traffic to its destination compartment in a synchronous wave. To facilitate the observation of the reporter, a fluorescent protein such as enhanced green fluorescent protein (EGFP) is usually fused to the SBP-bearing reporter. The transport kinetics varies depending on the cargo of interest. Most of the cargos reach the Golgi apparatus after 15 min of incubation with biotin and the plasma membrane starting from 30 min of treatment with biotin (see RUSH-synchronized transport of gp135/podocalyxin as an example, Fig. 1b). Once exocytic vesicles delivered their content at the cell surface, soluble proteins are released to the extracellular medium. Integral or membrane-attached proteins then diffuse in the plasma membrane, are internalized, or are recycled with different kinetics, based on their intrinsic diffusion properties and on their interacting partners. To be able to analyze local delivery of these diverse classes of cargos, we developed Selective Protein Immobilization (SPI) to be used in combination with the RUSH assay [4]. SPI is based on the extracellular capture of RUSH-synchronized proteins after their exocytosis, thanks to the presence on the coverslip of specific antibodies. The antibodies coated on the coverslip prior to cell seeding, in contact with the ventral surface of cells, will immobilize the RUSH cargo once exposed to the extracellular space after exocytosis (see Note 1). Because a lot of RUSH cargos available are fused to EGFP and because a large variety of anti-GFP antibodies is accessible, we developed and describe here SPI capturing GFP (see Notes 2–4 for tips to set up SPI, using antibodies directed to other proteins) (Fig. 2). This chapter describes how to perform SPI by coating coverslips with anti-GFP antibodies, how to express and induce the trafficking of the RUSH-synchronized cargo, and methods to detect local exocytosis using surface immunolabeling or real-time imaging in living cells (using total internal reflection microscopy or spinning disk microscopy).

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Fig. 1 Principle of the retention using selective hooks (RUSH) assay. (a) The RUSH system is two-state secretory assay, allowing synchronization of the trafficking of a cargo of interest (reporter). Retention of the reporter in the donor compartment is mediated by interaction between the streptavidin-binding peptide (SBP) fused to it and streptavidin fused to a protein stably resident in the donor compartment. Synchronous release of the reporter is induced by addition of biotin, which competes out the SBP. The reporter is then free to traffic to its target compartment. To facilitate the observation of the transport of the reporter, an EGFP (or another fluorescent protein) is fused to the reporter. (b) HeLa cells expressing RUSH-synchronized gp135/podocalyxin (Str-KDEL_SBP-EGFP-gp135) were seeded on non-coated glass coverslips. Cells were fixed after the indicated time of incubation with biotin. Pictures were acquired using an epifluorescence microscope. Scale bar: 10 μm

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Materials

2.1 Coating of Coverslips

1. Carbonate buffer 0.1 M, pH 9.5: Weight 8.40 g of NaHCO3 and 3.56 g of Na2CO3. Add water to a volume of 900 mL. Mix and adjust pH to 9.5. Make up to 1 L with water. Sterilize by filtration through a filter with pores of 0.2 μm of diameter. See also Note 5. Store at 4
C.

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Fig. 2 Principle of selective protein immobilization (SPI) combined to the RUSH assay. (a, b) Glass coverslips are coated with anti-GFP antibodies prior to cell seeding and transfection. Cells are then transfected to express a GFP-tagged RUSH cargo. After addition of biotin, the GFP-tagged RUSH cargo is released and exocytosed at the cell surface where it is immobilized by the anti-GFP antibodies present on the surface of the coverslip

2. Poly-L-lysine solution: Dilute poly-L-lysine in sterile water to a final concentration of 0.01%. 3. Anti-GFP antibody: A large source of anti-GFP antibodies is available from various providers. We set up the SPI assay using a homemade rabbit polyclonal antibody from the recombinant antibody platform of the Institut Curie. This anti-GFP antibody was purified and concentrated at 5 mg/mL. As an indication, homogeneous and efficient coating was obtained, using 0.05 mg/mL as final concentration of the anti-GFP antibody diluted in carbonate buffer. (See Notes 6–8 for tips about the homogeneity and the efficiency of the coating). 2.2 Cell Seeding, Transfection, and Release of the RUSHSynchronized Cargo

1. Glass coverslips. 2. Cell culture Petri dishes or multi-well plates. 3. Culture medium adapted to the cells of interest see Note 9. 4. EGFP-tagged RUSH cargo of interest (RUSH plasmids from the Perez lab are available from Addgene: https://www. addgene.org/Franck_Perez/). 5. Transfection reagent adapted to the cell line used. 6. A 4 mM biotin stock solution: Weight 48 mg of D-biotin (Sigma, ref. B4501), dissolve into 50 mL of culture medium. After complete dissolution, sterilize by filtration through a filter with pores of 0.2 μm of diameter. Store at 4
C for 3 months.

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1. Ice-cold phosphate buffer solution (PBS). 2. A 1% paraformaldehyde solution prepared in PBS. 3. Primary antibody: Either anti-GFP or anti-SBP antibodies. 4. Fluorochrome-conjugated secondary antibodies adapted to the primary antibody used. 5. Mounting medium: Weight 6 g of glycerol and 2.4 g of Mowiol 4–88. Add 6 mL of distilled water. Incubate for 2 h at room temperature under agitation. Add 12 mL of 0.2 M Tris–HCl pH 8.5. Agitate for 2 h at 50
C until it completely gets dissolved. Centrifuge for 15 min at 4000  g. Store aliquots at 20
C for several months.

2.4 Real-Time Imaging of Local Exocytosis by Total Internal Reflection Fluorescence or Spinning Disk Microscopy

1. Imaging medium such as Leibovitz’s medium is used. This medium does not need a high CO2 environment to image the cells during several hours. It is better to select Leibovitz without phenol red to reduce background. 2. Chambers adapted to glass coverslips or Chamlide magnetic chamber. 3. Inverted fluorescence microscope equipped with a thermostated incubation chamber, either spinning disk or TIRF microscope. 4. Immersion oil.

3

Methods

3.1 Coating of Coverslips with Anti-GFP Antibodies

1. Immerse glass coverslips in carbonate buffer 0.1 M pH 9.5. Incubate for 1 h at 37
C. 2. Transfer coverslips in poly-L-lysine 0.01% solution. Coverslips will float on the solution. Keep in mind that the poly-L-lysinecoated surface is the one facing the solution. Incubate for 1 h at 37
C. 3. Wash coverslips three times in PBS. Let the coverslips dry before proceeding to the next step. 4. Prepare 40 μL drops for 12 mm glass coverslips or 100 μL drops for 25 mm coverslips of anti-GFP solution, diluted in carbonate buffer on a parafilm foil. 5. Flip coverslips upside down with the face coated with poly-Llysine immersed in the antibody solution. 6. Add a humidified filter or tissue to create a humid environment and prevent dehydration. 7. Incubate for at least 3 h at 37
C (incubation might be prolonged overnight if more convenient). 8. Wash coverslips twice with PBS.

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9. Transfer the coated coverslips in a multi-well plate or a dish containing complete culture medium adapted for the cells of interest. 10. Seed the cells at the appropriate confluence for the transfection method of choice. Allow cell seeding for several hours or overnight before proceeding to transfection of the plasmid coding for the RUSH cargo. 3.2 Expression and Release of the Retention Using Selective Hooks-Synchronized Cargo

1. Transfect the cells to introduce the plasmid coding for the EGFP-tagged RUSH cargo. (See Note 10 for comments about transfection methods compatible with the RUSH assay). 2. From 18 h to 24 h post-transfection, replace medium containing transfection mix by pre-warmed complete medium containing 40 μM biotin. 3. Incubate for the desired time at 37
C. Trafficking kinetics is variable depending on the cargo. Consequently, it is critical to assess the trafficking kinetics of the cargo of interest prior to using the SPI assay. Proceed to the detection of local exocytosis either by surface immunolabeling or by real-time imaging in living cells. If real-time imaging is used, read Subheading 3.4, since the release of the cargo should be performed in chamber or dish adapted to the microscope to be used. Instead of using glass coverslips, some imaging dishes might be used, meaning that these dishes have to be coated with the anti-GFP antibodies.

3.3 Analysis of Local Exocytosis by Surface Immunolabeling

To visualize the exocytosed cargo only, an immunolabeling without permeabilization of the cells may be performed (Fig. 3). Antibodies directed to EGFP or to SBP might be used. Note that using antibodies from an animal species different from the one used for the coating of coverslips is critical. It also makes sense to use an antiEGFP antibody with a different epitope than the one used for coating to avoid any competition for binding to the exocytosed EGFP-tagged RUSH cargo. To perform surface immunolabeling, not fixing the cells is preferable to avoid the entry of the antibodies through small holes induced by fixation. 1. Following the steps described in the previous section, immediately wash coverslips with ice-cold PBS. Keeping the cells at 4
C is essential to stop trafficking of the cargo, preventing both further exocytosis of the cargo and endocytosis, which would lead to internalization of the antibody used for immunostaining. 2. Dilute the primary antibody (e.g., anti-EGFP or anti-SBP) in ice-cold PBS at the recommended concentration for immunofluorescence.

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Fig. 3 Local exocytosis of RUSH-synchronized CollagenX analyzed using surface immunolabeling. HeLa cells seeded on glass coverslips coated with rabbit polyclonal anti-GFP antibodies were transfected to express Str-KDEL_SBP-EGFP-ColX (first raw and green channel on the merged images). Biotin was added for 35 min, and then cells were processed for surface immunolabeling using mouse monoclonal anti-GFP antibodies (second raw [inverted colors] and red on merge). Scale bar: 10 μm

3. Prepare drops of the diluted primary antibody on a parafilm foil (or another type of holder) placed on ice. 25–50 μL and 100–150 μL drops will be sufficient for immunolabeling of cells seeded on 12 mm or 25 mm glass coverslips, respectively. 4. Incubate coverslips upside down on the drops for 45 min on ice. The cells are immersed in the antibody solution. 5. Wash three times with ice-cold PBS. 6. Fix with paraformaldehyde 1% for 15 min at room temperature. 7. Wash three times with PBS. The cells are fixed at this stage, and the rest of the protocol might be performed at room temperature.

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8. Dilute the fluorochrome-conjugated secondary antibody in PBS at the recommended concentration for immunofluorescence. 9. Prepare drops of the diluted secondary antibody on a parafilm foil (or another type of holder). 10. Incubate coverslips upside down on the drops for 30 min at room temperature. 11. Wash three times with PBS. 12. Mount the coverslip in a drop of Mowiol, deposited on a glass slide. The cells are immersed in the Mowiol between the coverslip and the slide. The mounting solution might also contain a nuclear dye such as Hoechst or DAPI (40 ,6-Diamidine-20 phenylindole dihydrochloride) to counterstain nuclei. 13. Allow the mounting medium to polymerize for at least 3 h prior to proceeding to observation with a fluorescence microscope. 3.4 Analysis of Local Exocytosis Using Real-Time Imaging in Living Cells

Real-time fluorescence microscopy in living cells allows monitoring dynamically the first events of exocytosis of the RUSHsynchronized cargo. This can be achieved using spinning disk microscopy (Fig. 4a) or using total internal reflection fluorescence (TIRF) microscopy (Fig. 4b). TIRF microscopy enables observation of the events occurring only at the ventral surface of cells and thus events occurring at the plasma membrane [5]. The protocol for imaging local exocytosis in real time is the same using either a spinning disk microscope or a TIRF microscope. 1. Follow the protocols described in Subheadings 3.1 and 3.2 until transfection. 2. After transfection, transfer the coverslips in a chamber adapted to the coverslips and the microscope stage. 3. Fill with medium adapted to video microscopy such as Leibovitz’s medium without phenol red. 4. Set up acquisition parameters (e.g., focal plane, wavelengths, exposure time, interval time between acquisitions, etc.). See Note 11 for advice to set up the TIRF angle. 5. Acquire three or more pictures. These pictures correspond to the retention state (absence of biotin) when no exocytic events are expected to occur. 6. Replace medium by pre-warmed imaging medium containing 40 μM biotin. Alternatively, imaging chamber might be filled directly with medium containing 40 μM biotin if the time taken to set up acquisition parameters is shorter than the time of

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Fig. 4 Local exocytosis of RUSH-synchronized CollagenX monitored using real-time imaging in living cells. HeLa cells transiently co-expressing RUSH-synchronized CollagenX (Str-KDEL_SBP-EGFP-ColX) and PaxillinmCherry were seeded on coverslips coated with rabbit polyclonal anti-GFP antibodies. Biotin was added at time zero. Transport of SBP-EGFP-ColX was analyzed using real-time imaging with a spinning disk microscope (a) or a TIRF microscope (b). Scale bars: 10 μm. (a) ©2019 Fourriere et al. (Originally published in The Journal of Cell Biology. https://doi.org/10.1083/jcb.201805002)

appearance of the first exocytic events. In any case, writing down the time of addition of biotin is essential to define time zero. 7. Resume acquisition. 8. Continue time-lapse acquisition as long as necessary to visualize exocytic events.

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Notes 1. The efficacy of SPI depends on the relative speed of immobilization (linked to the antibody-antigen affinity and to the concentration of the coated antibody) versus the relative speed of diffusion. For example, it is particularly efficient to analyze the exocytosis of the poorly diffusive Collagen X, while the signal is less contrasted when monitoring secretory soluble GFP [4]. 2. While using combination of the RUSH and SPI assays, make sure that the antibody chosen for the SPI assay is directed to the extracellular part of the protein of interest. Alternatively, if using an antibody directed against SBP or the fluorescent protein, make sure that these tags are fused to the luminal part of the protein of interest. 3. As mentioned earlier, we set up the SPI assay, using a homemade rabbit polyclonal anti-GFP antibody. However, this assay can be set up using other antibodies—either other anti-EGFP antibodies or antibodies directed to other targets. In a recent study [4], we also used anti-mCherry and anti-VSVG (Vesicular Stomatitis Virus G protein) antibodies for immobilization of mCherry-tagged RUSH cargo and RUSH-VSVG, respectively. If the antibody chosen is directed against your protein of interest, which is also endogenously expressed, the endogenous expression of the protein of interest might compete with the RUSH-synchronized exogenously expressed protein for capture. For this reason, using an antibody directed to the tag is an advantage. This also avoids performing multiple set-up experiments while changing antibodies. 4. The combination of the RUSH and SPI assays is also usable for multiple color imaging. In this case, two or more RUSH cargos tagged with different fluorescent proteins will be used. In consequence, the SPI assay, mixing two or more specific antibodies, has to be set up. 5. Carbonate buffer is quite easy to prepare. However, commercial enzyme-linked immunosorbent assay (ELISA) coating buffer can be used to treat coverslips and dilute the antibodies for SPI. 6. Homogeneity of the coating can be assessed by incubating the anti-GFP-coated coverslips with a fluorochrome-conjugated appropriate secondary antibody. Then use epifluorescence microscopy to observe coverslips. Focal plane might be difficult to set up. Fluorescence is expected to be homogenously distributed. If some zones are not fluorescent, bubbles might probably be present while incubating the coverslips during the coating procedure.

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7. Be careful to not scratch the coverslips while handling them with tweezers during the coating procedure and during their transfer to dishes or plates prior to cell seeding. 8. To check efficacy and saturation of the coating of coverslips with anti-GFP antibodies, coated coverslips can be incubated with serial dilutions of recombinant EGFP. Measure fluorescence intensity of the coverslip surface using an epifluorescence microscope. 9. One critical parameter for the use of the RUSH assay to synchronize the transport of cargos is to avoid excess of biotin in the culture medium (and/or serum). The presence of biotin prevents the efficient retention of the reporter. For example, Dulbecco’s Modified Eagle Medium (DMEM) GlutaMax from Thermo that we routinely use does not contain additional biotin and can be used efficiently with the RUSH assay. In contrast, we noticed that OptiMEM, DMEM-F12, and RPMI-1640 (Roswell Park Memorial Institute) from Thermo do contain biotin and prevent synchronization of the transport of the RUSH cargo. In these cases, low amount of avidin (or streptavidin) can be added in the medium to trap biotin contained in the medium and enable efficient retention. For example, we use 107 M (7 μg/mL) of avidin in OptiMEM and 0.4  108 M (0.28 μg/mL) in DMEM-F12 (both supplemented with 10% serum). In this condition, the addition of 40 μM biotin (final concentration) in the medium is still efficient to induce the release of the RUSH cargo. 10. We routinely use the calcium phosphate method to transfect HeLa cells [6]. Some transfection reagents such as Lipofectamine induce a very strong expression per cell, which often prevents protein trafficking to occur. The expression level of the cargo will also impact on its transport kinetics. Consequently, testing several transfection methods (calcium phosphate, JetPEI, JetPRIME, electroporation, etc.) might be useful to use the RUSH assay efficiently. Please note that lentiviral transduction is also usable. This is adapted, in particular, to establish cell lines stably expressing RUSH cargos and choosing medium-expressing population (see ref. 7 for a detailed protocol to establish a RUSH-stable cell line). 11. The co-expression of a fluorescently tagged protein stably localized at the plasma membrane (e.g., Myr-Palm-mCherry or paxillin-mCherry) may be useful to set up the TIRF angle, given that before the first exocytic events occurred, the RUSHtagged cargo is absent from the plasma membrane.

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References 1. Boncompain G, Divoux S, Gareil N, de Forges H, Lescure A, Latreche L, Mercanti V, Jollivet F, Raposo G, Perez F (2012) Synchronization of secretory protein traffic in populations of cells. Nat Methods 9(5):493–498. https:// doi.org/10.1038/nmeth.1928 2. Barrette-Ng IH, Wu SC, Tjia WM, Wong SL, Ng KK (2013) The structure of the SBP-Tagstreptavidin complex reveals a novel helical scaffold bridging binding pockets on separate subunits. Acta Crystallogr D Biol Crystallogr 69 (Pt 5):879–887. https://doi.org/10.1107/ S0907444913002576 3. Wilson DS, Keefe AD, Szostak JW (2001) The use of mRNA display to select high-affinity protein-binding peptides. Proc Natl Acad Sci U S A 98(7):3750–3755. https://doi.org/10.1073/ pnas.061028198 4. Fourriere L, Kasri A, Gareil N, Bardin S, Bousquet H, Pereira D, Perez F, Goud B,

Boncompain G, Miserey-Lenkei S (2019) RAB6 and microtubules restrict protein secretion to focal adhesions. J Cell Biol 218 (7):2215–2231. https://doi.org/10.1083/jcb. 201805002 5. Axelrod D (1981) Cell-substrate contacts illuminated by total internal reflection fluorescence. J Cell Biol 89(1):141–145. https://doi.org/10. 1083/jcb.89.1.141 6. Jordan M, Schallhorn A, Wurm FM (1996) Transfecting mammalian cells: optimization of critical parameters affecting calcium-phosphate precipitate formation. Nucleic Acids Res 24 (4):596–601 7. Boncompain G, Perez F (2013) Fluorescencebased analysis of trafficking in mammalian cells. Methods Cell Biol 118:179–194. https://doi. org/10.1016/B978-0-12-417164-0.00011-2

Chapter 18 Combining Single Molecule Super-Resolution Imaging Techniques to Unravel the Nanoscale Organization of the Presynapse Christopher Small, Ramon Martı´nez-Ma´rmol, Rumelo Amor, Frederic A. Meunier, and Merja Joensuu Abstract The fusion of synaptic vesicles with the plasma membrane underpins neurotransmission. A number of presynaptic proteins play a critical role in overcoming the energy barrier inherent to the fusion of the negatively charged vesicular and plasma membranes. Emerging concepts suggest that this process is hierarchical and dependent on rapid and transient reorganization of proteins in and out of small nanoclusters located in the active zones of nerve terminals. Examining the nanoscale organization of presynaptic molecules requires super-resolution microscopy to overcome the limits of conventional light microscopy. In this chapter, we describe three super-resolution techniques that allow for the examination of the nanoscale organization of proteins within live hippocampal nerve terminals. We used (1) single-particle tracking photoactivated localization microscopy (sptPALM) to resolve the mobility and clustering of syntaxin1A (STX1A), (2) universal Point Accumulation Imaging in Nanoscale Topography (uPAINT) to study the mobility of a pool of vesicular-associated membrane protein 2 (VAMP2) transiting on the plasma membrane, and (3) subdiffractional Tracking of Internalized Molecules (sdTIM) to track VAMP2-positive recycling synaptic vesicles in conjunction with Cholera Toxin subunit B (CTB), which has recently been shown to be trafficked retrogradely from the presynapse to the cell body via signaling endosomes. Key words Exocytosis, Endocytosis, Nanoclustering, SNARE , Presynapse, Synaptic vesicle, Single particle tracking, Nanobodies

1

Introduction

1.1 Neurotransmission

Neurotransmission is a fundamental process that drives information transfer between neurons. Neurotransmitter-filled synaptic vesicles (SVs) lie at the core of this neuronal communication. Following the arrival of an action potential and the ensued influx of calcium into the presynapse, SVs fuse with the plasma membrane at the active zone, thereby releasing neurotransmitters into the synaptic cleft. This process is called exocytosis and underpins communication

Florence Niedergang, Nicolas Vitale and Ste´phane Gasman (eds.), Exocytosis and Endocytosis: Methods and Protocols, Methods in Molecular Biology, vol. 2233, https://doi.org/10.1007/978-1-0716-1044-2_18, © Springer Science+Business Media, LLC, part of Springer Nature 2021

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between neurons [1, 2]. The fusion of SVs with the presynaptic plasma membrane is driven by a series of protein–protein and protein–lipid interactions. N-ethylmaleimide-sensitive-factor attachment receptor (SNARE) proteins form “zipper-like” complexes that prime SVs for fusion with the plasma membrane, leading to the two opposing membranes undergoing fusion. These proteins include the vesicle (v-) SNARE protein, vesicular-associated membrane protein 2 (VAMP2), and the target membrane (t-) SNARE proteins synaptosomal nerve–associated protein 25 (SNAP25) and syntaxin1A (STX1A) [1]. The fusion of SVs with the plasma membrane is triggered by calcium-sensitive vesicular synaptotagmin-1 (SYT1) binding to plasma membrane phospholipid phosphatidylinositol bisphosphate [3, 4, 5]. Following exocytic fusion, SV components are subsequently retrieved via invagination and vesicular pinching off from the plasma membrane, through a process called compensatory endocytosis. SNARE proteins are maintained under strict spatiotemporal control, in order to promote vesicular fusion. Presynaptic proteins and SVs are subjected to Brownian motion, driving an overall homogenization. For this reason, changes in the nanoscale organization of proteins must occur to maintain local protein concentration differences and to ensure that sets of molecules involved in a given function are in the right place at the right time. Synaptic proteins are therefore locally sequestered, to drive both exocytosis and endocytosis. Protein nanoclustering during exocytosis [1], and endocytosis [4, 5], has been studied in the context of multiple synaptic protein functions. Notably, we have previously shown that secretagogue stimulation increases the number of STX1A nanoclusters at the plasma membrane of neurosecretory (PC12) cells and at the motor nerve terminals of Drosophila melanogaster larvae [6]. Munc18-1, which coordinates the engagement of STX1A within the SNARE complex, also forms nanoclusters under basal conditions. In addition, the number of Munc18-1 molecules per cluster is reduced in response to stimulation [7]. A pool of SYT1 is also organized in clusters on the plasma membrane [8, 9]. The ability of presynaptic proteins to form dynamic nanoclusters is intrinsically linked to the SV cycle, that is, fusion and retrieval of SVs from the plasma membrane. Adding to the complexity of synaptic organization, several distinct populations of SV pools, which are defined by varied release probabilities, have been described. SVs have traditionally been divided into three pools: the readily releasable pool (SVs that are docked and primed at the plasma membrane and which constitute the initial release component following stimulation), the reserve pool (a low-mobility pool of vesicles that constitutes the majority of vesicles in nerve terminals that are only released upon high levels of stimulation), and the recycling pool (a relatively mobile pool of SVs that responds to

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stimulation) [10]. Additional SV pools have also been described: vesicle-associated proteins are incorporated into the plasma membrane, following exocytosis to form a transient “surface pool” [11], and the intersynaptic SVs that are transported between nerve terminals in axons form the so-called super-pool of vesicles [11, 12], which do not appear to respond to stimulation [13, 14]. Understanding how the dynamic organization of presynaptic proteins help shape the SV cycle is a complex question that requires the combination of different super-resolution microscopy techniques. In this chapter, we discuss the use of complementary superresolution methods that can be employed to resolve the nanoscale organization of proteins involved in exocytosis in conjunction with the mobility of various SV pools. For a summary of super-resolution techniques presented in this chapter, see Table 1. A visual description of the principles of single-molecule imaging is provided in Figs. 1, 2, and 3, respectively. 1.2 Resolving the Presynaptic Nanoscale Organization Using Super-Resolution Imaging

Nerve terminals are confined and densely packed structures (roughly 1 μm in diameter), which makes imaging single molecules at the presynapse challenging. Conventional microscopy is limited by the diffraction of light, which restricts the resolution of presynaptic structures to ~200 nm. In the crowded environment of the presynapse, fluorescence microscopy, using proteins tagged with GFP, mCherry, or other commonly used fluorophores, leads to a large overlap between fluorescent emitters, which greatly impacts resolution. Due to these limits, super-resolution single-molecule imaging has been employed to unravel the nanoscale organization of presynaptic proteins. To achieve high spatial precision, singlemolecule super-resolution microscopy requires low density of photoconvertible (or photoactivated) molecules, so that the individual emission profile of each fluorophore can be determined. This can be accomplished by (1) modifying the angle of illumination to excite only a subset of fluorophores, (2) stochastic photoconversion (or photoactivation) of fluorophores, and (3) sparse labeling: low concentrations of externally applied fluorescent ligands designed to tag a molecule of interest at the plasma membrane or upon endocytic recycling. The details of these three super-resolution approaches are discussed in the following sections.

1.3 Illumination of Neurons During Super-Resolution Imaging

An important step in preventing overlapping fluorescent emitters involves the use of Total Internal Reflection Fluorescence (TIRF) microscopy. In contrast to traditional epifluorescence, TIRF microscopy involves emission beams being sent to the sample at an angle, thereby illuminating only a thin bottom section of the cell. This allows for selective excitation of fluorophores within a restricted evanescence field (~200 nm), immediately adjacent to the glass bottom of the dish. TIRF microscopy is optimal for imaging of proteins adjacent to the plasma membrane (sptPALM) and on

Mobility state of presynaptic proteins (membrane-associated proteins, vesicular proteins)

Plasma membrane proteins

Recycling/super pool of SVs

uPAINT

sdTIM

Synaptic population

sptPALM

Superresolution technique Disadvantages

Allows for imaging of recycling synaptic vesicles Due to the inherent recycling of SV molecules, certain trajectories may originate from and determination of their heterogeneous locations other than SVs (see Note 10) mobility states Length of procedure, due to pulse chase, allows No exposure to UV light required imaging of only one neuron per dish Requires stimulation to study activitydependent internalization

Allows for imaging of transmembrane proteins Limited surface expression of certain vesicular proteins (see Note 8) with extracellular epitopes Endocytosis of protein of interest during Does not require stimulation of neurons imaging (see Note 9) No exposure to UV light required Only one neuron should be imaged per cell culture dish

Allows for imaging of intracellular proteins that Short trajectories due to rapid bleaching of mEos2 compared to organic dyes (e.g., associate with the plasma membrane or Atto647N) organelles Can be performed under a variety of conditions May require co-transfection with an additional marker to determine neuronal morphology (stimulated, non-stimulated, drug Phototoxicity due to UV exposure treatment) Optimized for multiple biological systems (in vivo imaging in Drosophila larvae, PC12 neurosecretory cells) Multiple neurons can be imaged per dish

Advantages

Table 1 Description of super-resolution techniques for live imaging

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Fig. 1 Schematic representation of the sptPALM super-resolution technique for live imaging of membrane-associated Syntaxin1A-mEos2 (STX1A-mEos2) in nerve terminals of cultured hippocampal neurons using TIRF illumination (the evanescent field propagating 100–200 nm from the glass-sample interface). The image shows STX1A-mEos2 moving on the presynaptic plasma membrane, adjacent to the postsynaptic membrane

the plasma membrane (uPAINT). However, neuronal cultures, and especially axons, often extend beyond this thin illumination layer and, therefore, a deeper penetration depth (up to ~500 nm) can also be achieved with oblique illumination (adjustment of the incidence angle of the laser illumination to slightly smaller than the critical angle required in TIRF). Oblique illumination is well suited for imaging recycling SVs (sdTIM). 1.4 Photoactivated Localization Microscopy to Study the Nanoscale Organization of Membrane-Associated Proteins

Super-resolution can be achieved through controlled, stochastic photoconversion of fluorophores, with the majority of molecules being held in a non-emissive state. sptPALM is a technique that allows for the imaging of fluorescently tagged proteins at high localization precision, involving the use of photoconvertible fluorophores (Fig. 1). One of the photoconvertible fluorophores commonly used in sptPALM is mEos2. Excitation with low-level UV light (405 nm) causes mEos2 to randomly photoconvert from a green-to-red emitting state. mEos2 can then be excited using a 561 nm laser. Thus, stochastic photoconversion of mEos2-tagged

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Fig. 2 Schematic representation of the uPAINT super-resolution technique for live imaging of VAMP2-pHluorin–bound Atto647N-nanobodies on the plasma membrane of live hippocampal neurons using TIRF illumination (the evanescent field propagating 100–200 nm from the glass-sample interface). The abbreviation NB refers to nanobodies. The image shows a presynaptic nerve terminal filled with VAMP2-pHluorin-positive SVs and, following high potassium stimulation, VAMP2-pHluorin is translocated onto the plasma membrane and subsequently recognized by anti-GFP Atto647N nanobodies

proteins allows the sequential detection of single molecules at low density conducive to high localization precision. mEos2 can be used to tag a variety of proteins, including cytosolic proteins that associate with the plasma membrane and vesicular cargoes. Here, we describe the use of the mEos2-tagged STX1A (STX1A-mEos2) to examine its nanoscale mobility as an example of the applications of sptPALM [6, 15]. 1.5 Tracking the Surface Pool of Synaptic Vesicle Proteins Following Exocytosis

uPAINT is a super-resolution method for single-molecule imaging of proteins on the plasma membrane, which involves external application of nanobodies conjugated to a highly fluorescent fluorophore (Atto565, Atto647N, or Cy3) to label a protein of interest on the plasma membrane [16] (Fig. 2). These nanobodies

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Fig. 3 Schematic representation of the sdTIM super-resolution technique for resolving recycling SVs containing VAMP2-pHluorin–bound Atto565-nanobodies and signaling endosomes containing Alexa647-CTB, in live hippocampal neurons, using oblique illumination (propagating up to 500 nm from the glasssample interface). The abbreviation NB refers to nanobodies. The image shows a presynaptic nerve terminal filled with VAMP2-pHluorin-positive SVs. VAMP2pHluorin is translocated onto the plasma membrane following high potassium stimulation-induced exocytosis, which is followed by activitydependent uptake of VAMP2-pHluorin-bound Atto565 nanobodies and Alexa647-CTB into recycling SVs and signaling endosomes

commonly target the GFP-tag (or pHluorin, a pH-sensitive GFP variant) of proteins, but nanobodies with specificity to endogenous proteins have also been used [17]. uPAINT was first developed as a tool to track endogenous α-amino-3-hydroxy-5-methyl-4-isoxazolepropionic acid (AMPA) glutamate receptors in the postsynapse and GFP-tagged glycolipids (GPI-GFP) in the COS7 cell line [17]. More recently, we have adapted uPAINT to assess the mobility of the SV surface pool on the plasma membrane [13]. In this experimental setup, imaging of the surface pool of vesicular cargoes involves overexpressing a vesicular protein tagged at its luminal

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domain with pHluorin [18]. Following stimulation with a high potassium buffer, vesicles fuse with the plasma membrane and the luminal pHluorin-tag is exposed to the synaptic cleft and becomes available for anti-GFP/pHluorin nanobody binding. This allows nanoscale detection and tracking of single molecules of SV proteins transiting on the plasma membrane. In this chapter, we demonstrate the use of pHluorin-tagged VAMP2 (VAMP2-pHluorin) to investigate the surface pool mobility of the vesicular cargo on the plasma membrane as an example of the uPAINT application (Fig. 2). Cultured hippocampal neurons overexpressing VAMP2-pHluorin are stimulated with a high potassium buffer in the presence of low concentrations of anti-GFP Atto647N-labelled nanobodies (Atto647N-nanobodies). Following SV fusion with the plasma membrane, anti-GFP Atto647Nnanobodies bind to the exposed pHluorin-tag on the plasma membrane, and protein mobility can be assessed by tracking the fluorescence of the Atto647N-nanobodies in the far-red channel using TIRF illumination. Moreover, the transition of the luminal pHluorin-tag from the acidic pH of the SV lumen to the neutral pH of the extracellular environment leads to unquenching of the fluorophore: VAMP2-pHluorin becomes fluorescent, allowing the identification of active release sites. Thus, the pHluorin moiety enables both the tracking of bound nanobodies at the nanoscale and the localization of release sites from hippocampal neurons, following stimulation. 1.6 Tracking Single Recycling Synaptic Vesicles

sdTIM is a single-molecule localization application that has allowed us to track single recycling SVs with high localization precision [13, 14]. The sdTIM technique builds on the uPAINT technique by introducing a pulse-chase step to induce both exocytosis and compensatory endocytosis of VAMP2-pHluorin–bound Atto647N-nanobodies into the pool of recycling SVs in live presynapses. Similar to uPAINT, neurons are transfected with VAMP2-pHluorin. Neurons are stimulated for 5 min with a high potassium buffer that contains a low concentration of Atto565- or Atto647N-labeled nanobodies. The corresponding pHluorin unquenching is monitored on the microscope. This pulse step triggers fusion of SVs, allowing nanobodies to access and bind to VAMP2-pHluorin. After a 5-min pulse, the neurons are chased (washed) in imaging buffer to remove excess high potassium buffer and unbound nanobodies. Subsequently, neurons are allowed to rest in the imaging buffer for 10 min. During this time, a corresponding decrease in pHluorin fluorescence should be monitored. This step allows VAMP2-pHluorin–bound nanobodies to be internalized and become incorporated into the recycling SV pool. Depending on the fluorophore labeling of the ligand, we can commonly reach a ~35–40 nm localization precision, which is slightly lower than the diameter of an SV (~40 nm) [19]. Therefore,

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the mobility of the VAMP2-pHluorin–bound nanobody is mainly dictated by the mobility of the recycling vesicle rather than the mobility of the protein within the vesicle itself. Here, we describe the use of VAMP2-pHluorin–bound antiGFP Atto565-nanobodies and Alexa647-tagged Cholera Toxin Subunit B (Alexa647-CTB) to study the mobility of recycling SVs and signaling endosomes, respectively, as an example of the simultaneous dual-color sdTIM application (Fig. 3). We recommend using organic far-red dyes (i.e., Atto647N) when performing super-resolution imaging, using one channel for optimal signal-tonoise ratio. 1.7 Dual-Color Super-Resolution Imaging of Synaptic Molecules

In this chapter, we have described different approaches for deciphering the nanoscale organization of molecules: sptPALM as a readout of protein mobility at the presynapse, uPAINT, and sdTIM for examining mobility of surface and recycling SV pools, respectively. These techniques can be combined, in order to determine whether or not two proteins localize and cluster together and how two proteins act in concert with one another. In general, the simultaneous detection of two separate proteins of interest can be achieved in these experimental setups either by the combined use of (1) photoconvertible molecules (i.e., mEos2 photoconversion with 405 nm and detection following 561 nm excitation) and far-red dyes (Alexa647- or Atto647N-tagged ligands imaged with 642 nm excitation) or (2) red dyes (Alexa565- or Atto565-tagged ligands) and far-red dyes (Alexa647- or Atto647N-tagged ligands). When choosing the appropriate combination of super-resolution imaging techniques, it is worth noting that the use of sptPALM allows single-molecule detection and tracking of membrane-associated proteins as well as organelle-bound molecules such as those found on SVs or autophagosomes, whereas uPAINT and sdTIM are used for examining protein mobility on the plasma membrane or upon internalization, respectively. Here, we first demonstrate the use of sptPALM in tandem with uPAINT to study the mobility of STX1A-mEos2 at the presynapse (mEos2-tag is located on the C-terminus of the STX1A facing the extracellular space) and VAMP2-pHluorin–bound Atto647N-nanobodies transiting on the extracellular side of the presynaptic plasma membrane, respectively. We then demonstrate the use of sdTIM for simultaneous dual-color imaging to track the recycling vesicle mobility of VAMP2-pHluorin–bound Atto565-nanobodies in conjunction with internalization of Alexa647-CTB, which binds to ganglioside GM1 on the plasma membrane prior to its internalization into signaling endosomes and retrograde trafficking [20].

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Materials

2.1 Transfection and Plasmid Constructs

1. Lipofectamine 2000 transfection reagent 2. Neurobasal media 3. STX1A-mEos2 [6] 4. VAMP2-pHluorin from J Rothman (Yale University, New Haven, CT) [18]

2.2 Imaging Buffers and Ligands

1. Imaging/low potassium buffer: 5.6 mM KCl, 0.5 mM ascorbic acid, 0.1% BSA, 15 mM HEPES, 5.6 mM D-Glucose, 145 mM NaCl, 0.5 mM MgCl2, and 2.2 mM CaCl2, at pH ¼ 7.4, 290–310 mOsm 2. Stimulation/ high potassium buffer: 56 mM KCl, 0.5 mM ascorbic acid, 0.1% BSA, 15 mM HEPES, 5.6 mM D-Glucose, 95 mM NaCl, 0.5 mM MgCl2, and 2.2 mM CaCl2, at pH ¼ 7.4, 290–310 mOsm 3. Anti-GFP Atto565/647N-labeled FluoTag-Q single-domain camelid antibodies (Synaptic Systems) 4. Cholera toxin subunit-B (recombinant) labeled with Alexa Fluor 647 (ThermoFisher Scientific) 5. TetraSpeck™ Microspheres, 0.1 μm, fluorescent blue/green/ orange/dark red (ThermoFisher Scientific)

2.3 Equipment for Imaging

2.4 Software for Single Particle Tracking

For live-cell super-resolution imaging, neurons are imaged at 37  C in TIRF or oblique illumination, using a Roper Scientific iLas2 Ring-TIRF microscope with a CFI Apo 100/1.49 N.A. oil-immersion objective (Nikon Instruments), two Evolve 512 Delta EMCCD cameras (Photometrics) mounted on a TwinCam LS Image Splitter (Cairn Research) for simultaneous dualchannel imaging, a Perfect Focus System (Nikon), an iLas2 doublelaser illuminator (Roper Scientific) for 360 TIRF illumination and 405 nm laser (100 mW, Vortran Laser Technology), 561 nm laser (150 mW, Cobolt Jive) and 642 nm laser (100 mW, Vortran). Image acquisition is performed using MetaMorph software (version 7.10.2, Molecular Devices). A TIRF-quality ultra-flat quadruple beam splitter (ZT405/488/561/647rpc; Chroma Technology) for distortion-free reflection of lasers and QUAD emission filter (simultaneous quadruple laser filter set for imaging blue, green, red and far-red; ZET405/488/561/640 m; Chroma) are required. TetraSpeck™ microspheres are used for dual-camera alignment and TIRF angle calibration. 1. PALMTracer 2, version 2.1.0.28228 [21, 22] 2. MetaMorph (Molecular Devices)

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Methods Single-particle tracking was carried out in E18 (embryonic day) rat hippocampal neurons cultured on poly-L-lysine-coated glassbottom dishes (29 mm) until DIV21-22 (days in vitro). For a detailed description of the hippocampal neuron dissections and neuronal culturing, see Joensuu and Martı´nez-Ma´rmol et al. [14].

3.1

Transfection

1. Transfection is carried from DIV14-15 (see Note 1). Calculate the appropriate volume of the plasmid (typically 1–2 μg of DNA per dish; however, this amount will vary depending on the plasmid). For sptPALM, co-transfection with GFP is recommended, to visualize neuronal morphology (see Note 2). For double transfections, we recommend using 1:1 ratio of each DNA plasmid. 2. Add total volume of DNA required to an Eppendorf tube (1.5 ml) containing 100 μl of neurobasal media per dish. Similarly, add lipofectamine (2 μg) to a second Eppendorf tube (1.5 ml) with 50–100 μl per dish. Commonly, a 1:1 ratio of DNA to lipofectamine is used. 3. Incubate both DNA and lipofectamine solution for 5 min at room temperature (i.e., 22–25  C), and then combine the solutions and incubate for additional 15–30 min at room temperature. 4. Collect the conditioned media from dishes (store the collected conditioning media at 37  C), leaving a small amount covering the central glass bottom to prevent neurons from drying out. Gently mix the lipofectamine-DNA mix and add the appropriate volume dropwise to the central glass bottom. Incubate the neurons for 2 h at 37  C with 5% CO2 atmosphere (the incubation time may vary depending on the plasmid being used) (see Note 3). 5. Following 2 h of incubation, use the stored conditioning media to wash and remove the lipofectamine-DNA mix and to restore the original media on the culture dishes. Conditioning media contains growth and survival factors, which facilitate neuronal survival, following lipofectamine treatment. In the case of inadequate volume of collected conditioned media, refer to Note 3. 6. Neurons are then further incubated in the conditioning media (at 37  C, 5% CO2) and typically a minimum of 24 h is required to allow expression of the transfected construct.

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Fig. 4 Calibration of the Roper Scientific iLas2 Ring-TIRF dual-camera alignment using multi-fluorescent TetraSpeck™ microspheres. Fluorescence following 642 nm and 491 nm excitation was captured using two Evolve 512 Delta EMCCD cameras. Panels show simultaneous dual-color imaging and misalignment of cameras (on left), the effect of stepwise xy-alignment (in the middle), and rotational alignment of cameras with respect to one another (on right). Magnified images from the boxed areas are shown in the bottom row. Scale bar is 10 μm for full field of view panels, and 4 μm for zoomed-in panels 3.2 Calibration of the Roper iLas2 Ring-TIRF Microscope

For single-particle tracking, the TIRF angle of lasers and the alignment of the two cameras on the Roper microscope must be calibrated before each experiment. This is especially important for simultaneous dual-color imaging. Add TetraSpeck™ microspheres (0.1 μm, at 1:1000 dilution in imaging buffer) to a glass-bottom dish (29 mm). These beads emit fluorescence in the blue/green/ red/far-red spectra and can therefore be used to align and angle the lasers (Fig. 4). With the EMCCD cameras switched off, ensure that the correct dichroic mirror and emission filters are installed. Mount the sample onto the stage of the Nikon microscope body and localize the fluorescent beads.

3.2.1 Alignment of Roper Scientific iLas2 Ring-Total Internal Reflection Fluorescence Microscope Cameras

1. The alignment of the lasers must be optimized before the calibration of the TIRF angle (the TIRF angle will need to be recalibrated each time the laser alignment is configured). 2. The two Photometrics Evolve 512 Delta EMCCD cameras are mounted on a TwinCam LS Image Splitter that allows fine

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xy-alignment of both the transmitted and reflected ports. With the two-camera images of the TetraSpeck beads overlaid in the MetaMorph Live View window, adjust the angle of the emission beam splitter, using the adjustment screws marked as “H” and “V” to achieve pixel-to-pixel alignment between the two channels. If required, rotate one camera relative to the other until the beads are aligned, including those located in the periphery (see Note 4). 3.2.2 Calibration of Total Internal Reflection Fluorescence Angle

1. In the iLas2 illumination system, the position of the laser excitation is controlled by galvanometer scanning mirrors. An even illumination of the sample in TIRF mode is achieved with an annular illumination pattern defined by four coordinates (north/south/east/west). The position of the scan mirrors must be calibrated for each filter cube. 2. Select the point mode from the Calibration tab in the iLas2 GUI and perform an individual calibration for each of the four coordinates.

3.3

sptPALM

Use DIV21-22 hippocampal neuronal cultures grown on glassbottom dishes and co-transfected with STX1A-mEos2 and soluble GFP (see Note 2). Note that the following section describes the use of sptPALM to image the mobility of plasma membrane–associated STX1A-mEos2 (refer to Subheading 3.6 for dual imaging). 1. Wash neurons with low potassium imaging buffer to remove traces of culture medium. 2. OPTIONAL: Add an appropriate concentration of TetraSpeck™ microspheres diluted in low-potassium imaging buffer on the glass-bottom dishes and incubate the dishes at 37  C for 10 min (see Note 5). 3. Localize transfected neurons (e.g., using 491 nm excitation for GFP co-transfection). 4. Switch the 405 and 561 nm lasers on. The 405 nm laser, which photoconverts the mEos2-fluorophores, is kept at low power to generate continuous stochastic photoconversions at a relatively low rate that prevent overlap of fluorescent single molecule emitters. The 561 nm laser excites the photoconverted molecules and is maintained at high intensity throughout acquisition. We use 20 ms exposure time (50 Hz), and typically acquire 16,000 frames (see Note 6).

3.4

uPAINT

Use DIV21-22 hippocampal neuronal cultures grown on glassbottom dishes and transfected with VAMP2-pHluorin. Note that the following section describes the use of uPAINT to image the plasma membrane surface pool of VAMP2-pHluorin–bound Atto647N-nanobodies (refer to Subheading 3.6 for dual imaging).

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1. Wash neurons with low-potassium imaging buffer to remove traces of culture medium. 2. OPTIONAL: Add an appropriate concentration of TetraSpeck™ microspheres diluted in low-potassium imaging buffer on the glass-bottom dishes and incubate the dishes at 37  C for 10 min (see Note 5). 3. Neurons are mounted on the Roper iLas2 microscope specimen stage at 37  C for imaging. Axons positive for VAMP2pHluorin are selected in the green channel. 4. Application of high-potassium buffer containing Atto647Nnanobodies (3.19 pg μl1) is used to stimulate neurons. However, acquisition can also be carried out with nanobodies in low-potassium buffer under non-stimulated conditions. Acquire 16,000 frames (in the far-red channel) immediately following stimulation at 20 ms exposure (50 Hz) (see Note 6). 3.5

sdTIM

Use DIV21-22 hippocampal neuronal cultures grown on glassbottom dishes and transfected with VAMP2-pHluorin. Note that the following section describes the use of sdTIM to image SV recycling using VAMP2-pHluorin–bound Atto647N-nanobodies (refer to Subheading 3.7 for simultaneous dual-color imaging). 1. Neurons are washed with low-potassium imaging buffer to remove traces of culture medium. 2. OPTIONAL: Add an appropriate concentration of TetraSpeck™ microspheres diluted in low-potassium imaging buffer on the glass-bottom dishes and incubate the dishes at 37  C for 10 min (see Note 5). 3. Glass-bottom dishes are placed on the imaging platform at the Roper iLas2 microscope operating at 37  C. Axons positive for VAMP2-pHluorin are selected in the green channel. 4. Stimulate the neurons for 5 min with high-potassium buffer (containing 3.19 pg μl1 of Atto647N-nanobodies). During this time, pHluorin levels can be recorded in the green channel to detect active presynapses. 5. Wash the neurons with imaging buffer (4–5 times, 2 ml). This step removes residual nanobodies and high potassium buffer from the dish. 6. Chase the neurons for 10 min at 37  C to allow internalization of the VAMP2-pHluorin–bound Atto647N-nanobodies. The pHluorin signal can be monitored to check for internalization of VAMP2-pHluorin post-stimulation. 7. Acquisitions are taken in the far-red channel at 20 ms exposure time typically for 16,000 frames.

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The first example of dual-color imaging presented in this chapter involves simultaneous acquisition of STX1A-mEos2 (sptPALM) and VAMP2-pHluorin labeled with Atto647N-nanobodies (uPAINT) (Fig. 5). Use VAMP2-pHluorin and STX1A-mEos2 co-transfected hippocampal neurons grown on glass-bottom dishes at DIV21-22. 1. Neurons are washed with low-potassium imaging buffer to remove traces of culture medium. 2. Add an appropriate concentration of TetraSpeck™ microspheres diluted in low-potassium imaging buffer on the glassbottom dishes and incubate the dishes at 37  C for 10 min (see Note 5). 3. At the time of imaging, ensure the EMCCD cameras are properly aligned and that the TIRF angle is calibrated accordingly. 4. Mount glass-bottom dishes on the specimen stage of the Roper iLas2 microscope and identify VAMP2-pHluorin-positive neurons using the green channel. 5. Switch to the 405/561 nm (wide-field) lasers to observe photoconversion of STX1A-mEos2 molecules. Activate dual cameras and 405/561/642 nm (wide-field) lasers. 6. Stimulate neurons with high-potassium buffer containing antiGFP Atto647N-nanobodies. Acquisitions for sptPALM and uPAINT are taken immediately at 20 ms exposure (typically for 16,000 frames) as described above. Alternatively, add nanobodies to the dish diluted in low-potassium imaging buffer for imaging under non-stimulatory conditions.

3.7 Dual-Color Imaging Using sdTIM

The second example of dual-color super-resolution imaging involves tracking of two endocytic markers, Alexa647-CTB, and VAMP2-pHluorin–bound Atto565-nanobodies to image and track the internalization of ligands in signaling endosomes and recycling SVs, respectively (Fig. 6). For imaging, use hippocampal neurons grown on glass-bottom dishes and transfected with VAMP2pHluorin (note that Alexa647-CTB is internalized by binding to its natural target on the plasma membrane and does not require overexpression of a target protein) (see Note 7). 1. At the time of imaging, ensure that the EMCCD cameras are aligned and the TIRF angle is calibrated. 2. Mature presynapses are typically imaged around DIV21-22, at which stage neurons are washed with low-potassium imaging buffer to remove traces of culture medium. 3. Add the appropriate concentration of TetraSpeck™ microspheres diluted in low-potassium imaging buffer on the glassbottom dishes and incubate the dishes at 37  C for 10 min. The use of microspheres is particularly important during

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Fig. 5 Simultaneous dual-color single-particle imaging and tracking of STX1A-mEos2 by sptPALM and resident pool of plasma membrane (PM) VAMP2-pHluorin–bound Atto647N-nanobodies by uPAINT in live hippocampal neurons (DIV21). Acquisition was performed for over 3 min (10,000 frames). (a) Wide-field image of an axon positive for VAMP2-pHluorin (imaged using 491 nm excitation). Scale bar ¼ 4 μm. (b) Detailed view of selected region of interest (dotted square) from (a) showing VAMP2-pHluorin epifluorescence. (c) Plotted STX1A-mEos2 trajectories (10,000 frames). (d) Plotted trajectories of VAMP2-pHluorin–bound Atto647Nnanobodies (on the PM) (10,000 frames). (e) Merged image of trajectories of single molecules of STX1AmEos2 and VAMP2-pHluorin–bound Atto647N-nanobodies. Scale bar ¼ 1 μm. (f) MSD (μm2) over time (s) and (g) frequency distribution of diffusion coefficient values (μm2 s1) of STX1A-mEos2 and VAMP2-pHluorin–bound Atto647N-nanobodies

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Fig. 6 Dual-color sdTIM-imaging of recycling SVs containing VAMP2-pHluorin–bound Atto565-nanobodies (Atto565-NB) and Alexa647-CTB vesicular carriers actively transported in hippocampal axons (DIV21) for over 5 min (16,000 frames). (a) Merged trajectory images of SVs containing internalized VAMP2-pHluorin–bound Atto565-nanobodies and signaling endosomes containing Alexa647-CTB. Scale bar ¼ 5 μm. (b) Zoomed-in view of the plotted trajectories. Scale bar ¼ 1 μm. (c) MSD (μm2) over time (s) and (d) frequency distribution of diffusion coefficient values (μm2 s1) of Alexa647-CTB and VAMP2-pHluorin–bound Atto565-nanobodies

simultaneous dual-color imaging to reduce issues caused by camera misalignment and facilitate colocalization of trajectories (see Note 5). 4. Mount the glass-bottom dish on the Roper iLas2 microscope specimen stage. 5. Identify neurons that are positive for pHluorin signal (491 nm excitation). 6. Stimulate neurons for 5 min with high-potassium buffer (containing 3.19 pg μl1 of Atto565-nanobodies and 50 ng ml1 Alexa647-CTB).

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7. Following stimulation, residual nanobodies and CTB are washed with imaging buffer (8–10 ml), and the neurons are incubated for 10 min (in total) in the imaging buffer. 8. After the 10-min incubation, activate the 561- and 642-nm lasers and start acquisition (20 ms exposure (50 Hz), typically for 16,000 frames) (see Note 6). 3.8 Image Processing and Analysis

Single-molecule localization is detected using a wavelet-based segmentation analysis and the resulting trajectories computed using a simulated annealing-based tracking algorithm in PALMTracer 2, a custom-written program package that operates in MetaMorph software. PALMTracer uses Gaussian fitting of emission spectra for high-precision two-dimensional (2D) localization of molecules [21, 22]. For quantification of single-molecule mobility within active nerve terminals, regions of interest are drawn around boutons with unquenched pHluorin fluorescence signal detected following high-potassium stimulation. Trajectories lasting at least eight frames are reconstructed and the mean square displacement (MSD) is computed for each trajectory. Spatial detection limit is set at 0.106 μm. The MSD is fitted by the equation MSD(t) ¼ a + 4Dt, where D is the diffusion coefficient, a is the y-intercept, and t is time. Mean-square displacement (μm2) is calculated and plotted over a 0.2 s time frame. The frequency distribution of the diffusion coefficient (Log10 of μm2 s1) is quantified.

3.9

The use of single-molecule super-resolution microscopy has greatly improved our understanding of the contributions made by synaptic proteins to facilitate neurotransmission [23]. In this chapter, we demonstrate the use of sptPALM, uPAINT, and sdTIM to examine the dynamics of synaptic proteins within different components of the presynapse. The great advantage of these techniques is that they can be performed in tandem. Here, we have shown the application of sptPALM to unravel the nanoscale organization of STX1AmEos2 in conjunction with uPAINT imaging of VAMP2-pHluorin on the plasma membrane. We also demonstrate the use of sdTIM on activity-dependent internalization of VAMP2-pHluorin and CTB in the recycling pool and signaling endosomes, respectively. These methods, along with uPAINT, can potentially be adapted to study a variety of proteins involved in exocytosis, endocytosis, and the formation of different SV pools. One aspect of the nanoscale organization of synapses that is yet to be addressed by the techniques presented in this chapter is whether certain proteins are targeted to specific SV pools [24]. Defining the “molecular signature” of different SV pools is of great interest [25]. In particular, due to the majority of SVs (80%) constituting the reserve pool of nerve terminals [10], sptPALM could be performed on specific vesicular proteins (e.g., Vesicular Glutamate Transporter-1 (vGLUT1)mEos2) to further our understanding of the nanoscale organization of reserve pool vesicles and its dynamics.

Conclusions

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Notes 1. Transfection of primary neurons. In our experience, transfecting hippocampal neurons at DIV14-17 yields a better transfection efficiency compared to transfecting neurons at a later time point. However, optimal transfection time may vary depending on the construct and the experimental aims. 2. Co-transfecting with a cytosolic marker to determine neuronal morphology. The emission signal of mEos2-tagged proteins is very faint and bleaches quickly when excited by the 491 nm laser. Therefore, it may be challenging to identify transfected neurons without some level of photobleaching. Thus, co-transfection with a cytosolic marker (with a fluorescent tag that does not interfere with the emission spectrum of the photoconverted mEos2) is helpful to identify and visualize neurons prior to photoconversion. For co-transfection, we recommend mixing the two plasmids together for 2–3 min prior to addition of neurobasal media. 3. Conditioned media storage. We recommend storing conditioned media in the primary culture incubator within a falcon tube, with the cap slightly loose, in order to ensure that the pH of the conditioned medium is constant due to continued exposure to 5% CO2. 4. Roper iLas2 Ring-TIRF microscope dual-camera alignment and region of interest selection. While it is important to keep the EMCCD (electron-multiplying charge-coupled device) cameras aligned for simultaneous dual-color imaging, occasional misalignment toward the periphery of the field of view may occur. For this reason, we recommend restricting the region of interest at the center of the field of view during acquisition, where the cameras should be optimally aligned. 5. Use of TetraSpeck™ microspheres for alignment and drift correction. An appropriate concentration of TetraSpeck beads should be used during acquisition. Should the concentration be too high, perform wash steps in imaging buffer to remove excess microspheres. Allow time for beads to settle at the bottom of the dish to prevent movement of beads during acquisition. 6. Acquisition settings in MetaMorph software for real-time single-particle imaging. Digitizer (20 MHz), gain (1, 1) EM gain (111), and frames to average (1). Settings apply to acquisitions for sptPALM, uPAINT, sdTIM, and simultaneous dualcolor imaging. 7. Detection of Alexa647-CTB and VAMP2-pHluorin/Atto565nanobodies. As shown in Fig. 6, Alexa647-CTB and Atto565nanobodies often bind to different populations of neurons,

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that is, Atto565-nanobodies bind to VAMP2-pHluorin positive neurons and Alexa647-CTB less so. 8. Low plasma membrane expression of a protein construct. Certain proteins are expressed at higher levels on the plasma membrane than others. Proteins with high surface expression (e.g., VAMP2-pHluorin) are easier to identify and image based on their pH-dependent unquenching of pHluorin. Vesicular cargoes such as vGLUT1 and SV protein 2A (SV2A) have low steady-state surface expression compared to others such as VAMP2 [26], and based on our experience, they are more challenging to image. This may be due to variance in copynumber of certain proteins per vesicle [9] or to different endocytic rates post-stimulation. 9. Endocytosis of molecules during uPAINT. The fluorescent emission of Atto647N-nanobodies prior to bleaching is rapid (in the order of milliseconds) compared to the time scale of endocytic events. uPAINT therefore relies on continuous labeling of pHluorin-tagged proteins with Atto647N-nanobodies on the plasma membrane that quickly bleach after they are excited. However, it is worth noting that nanobodies may enter the cells outside the region of imaging and may laterally enter the imaging field. Consequently, it may become challenging to distinguish between individual molecules that are internalized and molecules that remain on the plasma membrane during acquisition. Therefore, it is optimal to perform uPAINT imaging immediately following application of nanobodies to prevent acquisition of internalized molecules. 10. Establishing internalization rate of protein of interest for sdTIM. Performing sdTIM with a given synaptic protein requires the time of internalization to be established. We have previously shown that following exocytosis, VAMP2-pHluorin is internalized in recycling SVs within 10 min post-stimulation, which is detected as a decrease in its mobility, following internalization into recycling SVs [13]. We therefore recommend using HRP-tagged nanobodies and performing electron microscopy to determine the correct localization and internalization of other proteins of interest [14].

Acknowledgments The super-resolution imaging was carried out at the Queensland Brain Institute’s (QBI’s) Advanced Microscopy Facility. We thank the Clem Jones Centre for Ageing Dementia Research (CJCADR) for its support. This work was also supported by an Australian Research Council (ARC) Discovery Project Grant (DP190100647), ARC Linkage Infrastructure, Equipment, and

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Facilities Grant (LE130100078) and The National Health and Medical Research Council (NHMRC) Senior Research Fellowship to F.A.M. M.J. is supported by an ARC Discovery Early Career Researcher Award (DE190100565) and The University of Queensland Early Career Researcher Grant (UQECR2057309). R.M.M. is supported by the Clem and Jones Foundation, the State Government of Queensland, and the NHMRC Boosting Dementia Research Initiative. C.S. is supported by the Research Training Program (RTP) Scholarship and a QBI top-up Scholarship. F.A.M. is an NHMRC Senior Research Fellow (1155794). CS and MJ performed the dissections and culturing of the primary neurons. CS, RMM and MJ performed the sptPALM and uPAINT simultaneous dual-color imaging, and MJ performed the simultaneous dual-color sdTIM experiments. CS, RA, RMM and MJ performed the dual-camera alignment. All authors wrote and edited the manuscript. FAM and MJ supervised the experiments. The authors of this chapter declare no conflict of interest. References 1. Padmanabhan P, Bademosi AT, Kasula R, Lauwers E, Verstreken P, Meunier FA (2019) Need for speed: super-resolving the dynamic nanoclustering of syntaxin-1 at exocytic fusion sites. Neuropharmacology 169:107554 2. Li P, Bademosi AT, Luo J, Meunier FA (2018) Actin remodeling in regulated exocytosis: toward a mesoscopic view. Trends Cell Biol 28(9):685–697 3. Stein A, Radhakrishnan A, Riedel D, Fasshauer D, Jahn R (2007) Synaptotagmin activates membrane fusion through a Ca2+dependent trans interaction with phospholipids. Nat Struct Mol Biol 14(10):904 4. Mund M, van der Beek JA, Deschamps J, Dmitrieff S, Hoess P, Monster JL, Picco A, Ne´de´lec F, Kaksonen M, Ries J (2018) Systematic nanoscale analysis of endocytosis links efficient vesicle formation to patterned actin nucleation. Cell 174(4):884–896. e817 5. Kaksonen M, Roux A (2018) Mechanisms of clathrin-mediated endocytosis. Nat Rev Mol Cell Biol 19(5):313 6. Bademosi AT, Lauwers E, Padmanabhan P, Odierna L, Chai YJ, Papadopulos A, Goodhill GJ, Verstreken P, Van Swinderen B, Meunier FA (2016) In vivo single-molecule imaging of syntaxin1A reveals polyphosphoinositide-and activity-dependent trapping in presynaptic nanoclusters. Nat Commun 7:13660 7. Kasula R, Chai YJ, Bademosi AT, Harper CB, Gormal RS, Morrow IC, Hosy E, Collins BM, Choquet D, Papadopulos A (2016) The

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Chapter 19 Induction of Ca2+-Dependent Exocytotic Processes by Laser Ablation of Endothelial Cells Arsila P. K. Ashraf, Sophia N. Koerdt, Nikita Raj, and Volker Gerke Abstract Ca2+ regulates a variety of cellular processes that are essential to maintain cell integrity and function. Different methods have been used to study these processes by increasing intracellular Ca2+ levels. Here, we describe a protocol to initiate Ca2+-dependent membrane-related events, using laser ablation by nearinfrared irradiation. This creates a rupture in the plasma membrane that allows the extracellular Ca2+ to enter the cell and thereby induce a receptor-independent Ca2+ increase. We report laser ablation protocols to study two different Ca2+-induced processes in human endothelial cells—membrane resealing and exocytosis of secretory granules called Weibel-Palade bodies (WPBs). Thus, laser ablation represents a technique that permits the analysis of different Ca2+-regulated processes at high spatiotemporal resolution in a controlled manner. Key words Ca2+, Laser ablation, Plasma membrane, HUVEC, FM4-64, WPB exocytosis

1

Introduction Increases in intracellular cytosolic Ca2+ concentration initiate diverse processes within a living cell, which affect cell motility, shape, and fate but can also trigger Ca2+-regulated membrane trafficking events and even cell death [1, 2]. Furthermore, Ca2+ is known to play a pivotal role in plasma membrane resealing in diverse tissues and cell types [3]. Aspects of Ca2+-dependent cellular processes have been studied extensively, but precise spatiotemporal information of these processes, necessary to improve our understanding of them, is still lacking in many cases. Cytosolic Ca2+ elevation can be achieved experimentally in a number of different ways. For instance, classical studies employ receptor-based Ca2+ stimulation, where an agonist binds to its receptor on the cell membrane [4], which, in turn, elicits a chain of reactions that culminate in the increase of intracellular Ca2+ concentration. Here, Ca2+ entering through plasma membrane–

Florence Niedergang, Nicolas Vitale and Ste´phane Gasman (eds.), Exocytosis and Endocytosis: Methods and Protocols, Methods in Molecular Biology, vol. 2233, https://doi.org/10.1007/978-1-0716-1044-2_19, © Springer Science+Business Media, LLC, part of Springer Nature 2021

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resident ion channels and/or depletion of intracellular Ca2+ stores (like the endoplasmic reticulum) contribute to the rise in cytosolic Ca2+ levels [5]. In this chapter, we describe a technique to induce cytosolic Ca2+ elevation in a non-receptor based manner. In a live cell system, laser ablation of the plasma membrane by near-infrared irradiation induces Ca2+ increase by generating a rupture or wound in the cell membrane [6, 7]. This rupture makes way for the entry of Ca2+ from the extracellular medium. Connecting this setup to an imaging system allows the monitoring of processes initiated by the Ca2+ influx with high spatial and temporal resolution. Thus, this method of laser ablation can be used to study both Ca2+-initiated resealing of ruptured plasma membrane areas and exocytosis of secretory granules. To visualize the rupture of the membrane, a membraneimpermeable lipid dye like FM4-64 [8] is used (or alternatively, similar styryl dyes with different spectral characteristics, e.g., FM1-43) [9]. FM4-64 binds to the outer leaflet of the membrane, yielding a weak fluorescence signal. Upon laser ablation, dye present in the buffer rapidly enters the ruptured cell through the hole generated in the plasma membrane (Fig. 1a, b). Consequently, the dye can access and bind to intracellular membranes, which results in a significantly increased fluorescence signal (Fig. 1c). Dye uptake continues until the rupture is resealed, usually within tens of seconds (Fig. 1c). The FM4-64 uptake assay is thus a useful readout of the membrane resealing capability of a cell [6]. Cytosolic Ca2+ elevation by cell membrane wounding can also elicit the exocytosis of secretory granules, such as Weibel-Palade bodies (WPBs), found in vascular endothelial cells. WPBs serve as storage granules for factors controlling vascular homeostasis, for example, the coagulant glycoprotein von Willebrand factor (VWF) and the leukocyte receptor P-selectin [10–12]. Typically, WPB exocytosis is triggered by agonists that elevate cytosolic Ca2+ levels through receptor-mediated pathways. Importantly, mechanical ruptures of the endothelial plasma membrane, caused, for instance, by mechanical wear and tear or blood vessel injury, also induce Ca2+-dependent WPB secretion [13, 14]. WPB exocytosis can be

Fig. 1 Laser ablation assay. (a) A single cell in medium containing 2.5 mM Ca2+-supplemented Tyrode’s buffer and membrane-impermeable FM4-64 dye is irradiated with a near-infrared laser at its plasma membrane. (b) Ca2+ and FM4-64 enter into the cell through the ablated area. (c) FM4-64 binds to intracellular membranes, resulting in an increased fluorescent signal. FM4-64 uptake continues until the ablated area is resealed by Ca2+-activated mechanisms

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monitored by ectopically expressing labeled WPB cargo, such as VWF-GFP in endothelial cells. Individual WPB-plasma membrane fusion events can then be visualized in live-cell microscopy by the sharp increase of GFP fluorescence signal that occurs upon neutralization of the acidic intraluminal pH of WPBs [15]. In summary, laser ablation is a convenient technique to analyze 2+ Ca -stimulated cellular processes, which allows for spatiotemporal stimulus control, and which is independent of receptor agonists or Ca2+ ionophores. The method described here for endothelial cells is sufficiently flexible that it could be extended to other systems to shed light onto the fundamental aspects of critical Ca2+-regulated processes.

2

Materials

2.1 Cell Culture and Transfection

1. Human Umbilical Vein Endothelial Cells (HUVEC): Cells are isolated from umbilical cords according to ref. 16 or obtained from commercial sources. 2. HUVEC growth medium: Endothelial cell growth medium (ECGM-2) and Earle’s Medium M-199 are mixed at a ratio of 1:1. ECGM-2 is supplemented with 30 μg/mL gentamicin, 15 ng/mL amphotericin B, and 100 I.U. heparin. M199 Medium contains 10% fetal bovine serum superior, 30 μg/ mL gentamicin, 15 ng/mL amphotericin B, and 100 I.E. heparin. Store all media in sterile bottles at 4
C. 3. Dulbecco’s phosphate-buffered saline (DPBS). 4. Trypsin/EDTA solution (0.05%/0.02%). 5. EVOS fl digital inverted fluorescence microscope. 6. Fifteen-milliliter Falcon tubes. 7. Centrifuge (to spin 200  g for 4 min at room temperature). 8. Plasmids for recording exocytosis: VWF-GFP, as described in ref. 17 (kindly provided by Tom Carter (St. George University, London, UK)). 9. Electroporation cuvettes, 2 mm. 10. Transfection reagent: Amaxa HUVEC Nucleofector Kit-OLD or self-made buffer (4 mM KCl, 10 mM MgCl2, 10 mM sodium succinate, 100 mM NaH2PO4, pH 7.4). Store at 4
C in a sterile condition. 11. Nucleofector™ AAD 1001 Device. 12. Coating reagent: 50 μg/mL rat-tail collagen I in 0.02 M acetic acid. Store at 4
C in a sterile bottle. 13. μ-Slide eight-well glass-bottom dishes. 14. 60 mm-sized cell culture dishes.

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2.2 Laser Ablation Assay

1. Imaging medium: Mixed endothelial growth medium supplemented with 20 mM HEPES, pH 7.2. Store at 4
C. 2. Tyrode’s Buffer [18]: 140 mM NaCl, 5 mM KCl, 1 mM MgCl2, 10 mM Glucose, 10 mM HEPES, pH 7.2, and supplemented with either 2.5 mM Ca2+ or 100 μM EGTA. Sterile filter and store at 4
C. 3. FM4-64 stored at 20
C. 4. Immersion oil for objectives for 37
C observations.

2.3 Image Acquisition: Live Cell Confocal Microscopy

1. Laser scanning confocal microscope equipped with multichannel photomultiplier GaAsP detector and standard photomultiplier tube detector. 2. Incubator connected with CO2. 3. Plan-Apochromat 63/1.4 oil immersion DIC objective. 4. Excitation laser 488 nm (LASOS Argon laser). 5. Confocal software for image acquisition adapted to the microscope. 6. External Chameleon Vision Tunable Non-linear Optics (NLO) laser.

2.4 Software for Image Analysis

3

Image analysis is performed with Fiji on ImageJ [19].

Method Carry out all the experiments in sterile conditions under a biosafety hood and using reagents pre-warmed to 37
C, unless mentioned otherwise.

3.1 Human Umbilical Vein Endothelial Cell Culture 3.1.1 Human Umbilical Vein Endothelial Cell Culture for FM4-64 Uptake Assay

1. Culture HUVECs in 60 mm dishes with 4 mL mixed HUVEC growth medium at 37
C and 5% CO2. For expansion, split cells at a maximum ratio of 1:3. Use nearly confluent cells at passages 2–3 for the experiment. 2. Coat a μ-slide eight-well glass-bottom dish with 200 μL coating reagent per well and incubate at 37
C for 20–30 min (see Notes 1 and 2). 3. Wash each well of the eight-well dish twice with DPBS. 4. Add 200 μL of HUVEC growth medium to each well of the eight-well dish and store in 37
C, 5% CO2 incubator until further use. 5. From a nearly confluent 60 mm HUVEC culture dish (90–95% confluency; approximate culture surface 21 cm2), aspirate the medium and wash the cells with 1 mL DPBS (see Note 3).

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6. Aspirate the DPBS, add 1 mL Trypsin/EDTA, and incubate at 37
C, 5% CO2 for 1–2 min. 7. Check the cells under EVOS microscope to see if they are detached from the dish surface (see Note 4). 8. Add 3 mL of HUVEC growth medium to the trypsin-cell mixture and ensure that the contents are mixed well. Transfer to a 15 mL Falcon tube (see Note 4). 9. Centrifuge the cells at 200  g for 4 min at room temperature. 10. Discard the supernatant and resuspend the pellet in 1 mL of HUVEC growth medium. 11. Take the eight-well dish out of the incubator and add 50 μL of the cell suspension to each well of the dish (approximately 1 cm2 surface area per well). 12. Distribute the cells evenly by shaking the eight-well dish. Incubate the dish at 37
C, 5% CO2 for 24 h. 3.1.2 Human Umbilical Vein Endothelial Cell Transfection for Assaying Exocytosis

1. Prepare the electroporation cuvettes by adding the corresponding plasmid DNA, i.e. VWF-GFP (4 μg), for transfection per 60 mm culture dish (see Note 5). 2. Passage HUVECs as mentioned in Subheading 3.1.1 until step 10. 3. Discard the supernatant and resuspend the pelleted cells in 100 μL of transfection reagent. 4. Transfer 95 μL of the above cell suspension into the electroporation cuvette and mix well by gently vortexing. Get rid of air bubbles by tapping the cuvette on a hard surface. 5. Place the cuvette in the Nucleofector™ device and electroporate the cells using the Amaxa nucelofector program U-001. 6. Immediately, transfer the whole suspension to an Eppendorf vial filled with 1 mL HUVEC growth medium (see Note 6). It is helpful to use some of the medium to flush the suspension form the cuvette. 7. Transfer 50 μL of the transfected cell mixture to each well of the eight-well dish already containing medium. 8. Distribute the cells evenly and incubate the dish at 37
C, 5% CO2 for 24 h.

3.2 Preparation of Human Umbilical Vein Endothelial Cells for Laser Ablation Assay

1. Check cell confluency after 24 h in the eight-well slide under the EVOS microscope. If the cells are sub-confluent (50–70% confluency), proceed for live-cell laser ablation assay (see Note 7). 2. Aspirate the medium from each well of the eight-well dish and add 200 μL of imaging medium.

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3. Wash the well selected for imaging once with Tyrode’s buffer and add 240 μL Tyrode’s buffer (with Ca2+ or EGTA). 4. Add 6 μL of 200 μg/mL FM4-64 dye (final concentration: 5 μg/mL) and mix well with the buffer. 5. Immediately, take the dish to the microscope incubation chamber pre-warmed to 37
C. 3.3

Microscopy

1. Pre-warm the microscope incubation chamber to 37
C to ensure viability of the cells during imaging. 2. Open the ZEN 2.1 SP3 software on the computer and start with the NLO hardware configuration database. 3. Set up the ablation protocol on ZEN 2.1 SP3 software: (a) Select the wavelength of Chameleon laser as 820 nm (see Note 8) on the “Bleaching” tab, and set the laser power to 16% (see Note 9). (b) Set bleaching for two iterations with a pixel dwell of 77 μs in “zoom bleach” mode. (c) Set the “start bleaching” after two scans, so that two unbleached frames are obtained for the subsequent analysis. (d) Checkmark box “Safe bleach for GaAsP detector.” (e) For the live cell recording, set 100 cycles with no intervals on the “Time Series” tab. 4. Add a drop of 37
C immersion oil onto the objective and mount the eight-well dish on the microscope stage (see Note 10).

3.4 Laser Ablation and Confocal Image Acquisition

3.4.1 FM4-64 Uptake Assay

We use the “channel mode” imaging setup with one track. FM4-64 is excited by the 488 nm laser. Additionally, other fluorophores can be imaged together with FM4-64, using the appropriate lasers. For the acquisition, “Frame” scan mode is used with the frame size optimally adjusted to the zoom size. 1. Select FM4-64 as the dye on the “Imaging Setup” tab. 2. Set the pinhole to 1 Airy Unit (A.U.) and the 488 nm laser power to 2% (see Note 11) on the “Channels” tab. In the “Integration” mode, set the master gain in the range of 750–800 and the digital gain to 1.0. 3. Select the well with FM4-64 in Tyrode’s buffer and adjust the focus in the “live” mode (see Note 12). 4. Search for cells, which have an even FM4-64 staining of their membrane (see Notes 13 and 14). 5. Focus on the basal plane of a single cell, where FM4-64 intensity is optimal (see Note 15).

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Fig. 2 FM4-64 uptake assay. HUVECs were ablated in the presence of 5 μg/mL FM4-64 in Tyrode’s buffer supplemented with either 2.5 mM Ca2+ or 100 μM EGTA. The red triangles indicate the area of laser irradiation (wound size 2 μm2). In the presence of Ca2+, FM4-64 fluorescence is limited to the ablated area. FM4-64 uptake is more pronounced and sustained when the extracellular milieu lacks Ca2+. Scale bars ¼ 10 μm

6. Take a snap of the cell and draw a circular region of interest (ROI) of 20 pixels (see Note 16) in diameter (corresponds to 2 μm2 surface area) on the flat edge (see Notes 15 and 17). 7. Optional: If required, “continuous” mode.

adjust

the

focus

in

the

8. Start the ablation protocol. Avoid any type of external disturbances while the acquisition is progressing. A time series of 100 frames is recorded with the ablation occurring after the second frame. 9. Repeat steps 4–8 for more cell replicates in the well (see Note 18) and for different experimental conditions (see Note 19). 10. Save the acquired images for analysis (Fig. 2).

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3.4.2 Weibel-Palade Body Exocytosis Assay

1. Select both FM4-64 and GFP as the dyes on the “Imaging Setup” tab. 2. Set the pinhole to 1 Airy Unit (AU) and the 488 nm laser power to 2% (see Note 11) on the “Channels” tab. In the “Integration” mode, set the master gain in the range of 750–800 and the digital gain to 1.0. 3. Select the well with FM4-64 in Tyrode’s buffer and adjust the focus in the “live” mode (see Note 12). 4. Search for cells, which have an even FM4-64 staining of their membrane (see Notes 13 and 14) and an optimal expression of VWF-GFP.

Fig. 3 WPB exocytosis triggered by laser ablation. HUVECs transfected with VWF-GFP were subjected to laser ablation in Ca2+-containing medium in the presence of FM4-64 and observed by fluorescence microscopy. The red triangles indicate the area of laser irradiation (wound size 2 μm2). Exocytosis of WPB triggers an increase in VWF-GFP fluorescence intensity (due to neutralization of the acidic luminal pH of WPB upon fusion) and more rounded morphology of the VWF-GFP signal (marked by white arrows). FM4-64 restriction to the ablated area site shows successful resealing. Scale bars ¼ 10 μm

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5. Focus on the basal plane of a single cell, where FM4-64 intensity and VWF-GFP expression are optimal (see Note 15). 6. Take a snap of the cell and draw a circular ROI of 20 pixels (see Note 16) in diameter (corresponds to 2 μm2 surface area) on the flat edge of the cell (see Notes 15 and 17). 7. Optional: Adjust the focus plane in the “continuous” mode. 8. Start the ablation protocol. Avoid any type of external disturbances while the acquisition is progressing. A time series of 100 frames is recorded, with the ablation occurring after the second frame. 9. Repeat steps 4–8 for more cell replicates in the well (see Note 18) and for different experimental conditions (see Note 19). 10. Save the acquired images for analysis (Fig. 3). 3.5

Analysis

3.5.1 FM4-64 Uptake Assay

The uptake of FM4-64 is measured using “Plot Z-axis Profile” function in Fiji. A macro has been developed to automate the analysis for different cells (see later). In a nutshell, 1. Create a maximum-intensity projection of the acquired xyt image stack (see Notes 20 and 21). Draw the outline of the ablated cell of interest (COI) and select a part of a non-ablated cell for background (BG) correction (see Note 22). 2. Subtract the first image of the FM4-64 channel from all other images in this channel within the xyt stack. 3. Assess the fluorescence intensity changes for both COI and BG using the “Plot Z-axis Profile” function. 4. Process the resulting table of intensity values by subtracting BG from COI intensity at each time point. 5. Average the corrected fluorescence intensities obtained for all the time points and within each experimental condition. 6. Plot the average fluorescence intensities against time to obtain the final result, together with a measure of variance of your choice for each time point (Fig. 4).

Fiji Macro for FM4-64 Uptake Analysis

run("Close All"); run("Options...", "iterations=1 count=1 black edm=Overwrite"); // Choose a directory that contains nothing but the source images dir = getDirectory("Choose wisely..."); list = getFileList(dir); resultPath = dir + File.separator + "Results"

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Arsila P. K. Ashraf et al. File.makeDirectory(resultPath); fileWithoutEnding = ""; for (i=0; i