The Extracellular Matrix: Methods and Protocols [1st ed.] 978-1-4939-9132-7, 978-1-4939-9133-4

Table of contents :
Front Matter ....Pages i-xii
The Complex Interplay Between Extracellular Matrix and Cells in Tissues (Dimitra Manou, Ilaria Caon, Panagiotis Bouris, Irene-Eva Triantaphyllidou, Cristina Giaroni, Alberto Passi et al.)....Pages 1-20
Method for Studying ECM Expression: In Situ RT-PCR (Elena Caravà, Cristiana Marcozzi, Barbara Bartolini, Marcella Reguzzoni, Paola Moretto, Ilaria Caon et al.)....Pages 21-31
Visualizing the Supramolecular Assembly of Collagen (Mario Raspanti, Marcella Reguzzoni, Petra Rita Basso, Marina Protasoni, Désirée Martini)....Pages 33-44
Steady-State and Pulse-Chase Analyses of Fibrillar Collagen (Antonella Forlino, Francesca Tonelli, Roberta Besio)....Pages 45-53
Methods for Isolation and Characterization of Sulfated Glycosaminoglycans from Marine Invertebrates (Mariana P. Stelling, Ananda A. de Bento, Philippe Caloba, Eduardo Vilanova, Mauro S. G. Pavão)....Pages 55-70
Analysis of Proteoglycan Synthesis and Secretion in Cell Culture Systems (Chiara Paganini, Rossella Costantini, Antonio Rossi)....Pages 71-80
Analysis of UDP-Sugars from Cultured Cells and Small Tissue Samples (Sanna Oikari, Markku I. Tammi)....Pages 81-89
Methods for Hyaluronan Molecular Mass Determination by Agarose Gel Electrophoresis (Mary K. Cowman)....Pages 91-102
Hyaluronan Pericellular Matrix: Particle Exclusion Assay (Melanie A. Simpson)....Pages 103-110
Determination of Cell-Surface Hyaluronan Through Flow Cytometry (Daiana L. Vitale, Fiorella M. Spinelli, Laura Alaniz)....Pages 111-116
Method for Detecting Hyaluronan in Isolated Myenteric Plexus Ganglia of Adult Rat Small Intestine (Michela Bistoletti, Paola Moretto, Cristina Giaroni)....Pages 117-125
Analyzing Hyaluronidases in Biological Fluids (Christos Velesiotis, Stella Vasileiou, Demitrios H. Vynios)....Pages 127-142
Method for Studying the Physical Effect of Extracellular Matrix on Voltage-Dependent Ion Channel Gating (Eleonora Solari, Andrea Moriondo)....Pages 143-156
Methods for Monitoring Matrix-Induced Autophagy (Carolyn Chen, Aastha Kapoor, Renato V. Iozzo)....Pages 157-191
Method for Determining Gelatinolytic Activity in Tissue: In Situ Gelatin Zymography (Elin Hadler-Olsen, Jan-Olof Winberg)....Pages 193-199
Method for Determining Gelatinolytic Activity in Tissue Extracts: Real-Time Gelatin Zymography (Elin Hadler-Olsen, Jan-Olof Winberg)....Pages 201-210
In Vitro Spheroid Sprouting Assay of Angiogenesis (Fatema Tuz Zahra, Efrossini Choleva, Md Sanaullah Sajib, Evangelia Papadimitriou, Constantinos M. Mikelis)....Pages 211-218
Matrigel Plug Assay for In Vivo Evaluation of Angiogenesis (Pinelopi Kastana, Fatema Tuz Zahra, Despoina Ntenekou, Stamatiki Katraki-Pavlou, Dimitris Beis, Michail S. Lionakis et al.)....Pages 219-232
Exosomes from Cell Culture-Conditioned Medium: Isolation by Ultracentrifugation and Characterization (Anurag Purushothaman)....Pages 233-244
Preparation and Characterization of Tissue Surrogates Rich in Extracellular Matrix Using the Principles of Macromolecular Crowding (Adrian Djalali-Cuevas, Sergio Garnica-Galvez, Andrea Rampin, Diana Gaspar, Ioannis Skoufos, Athina Tzora et al.)....Pages 245-259
Novel Approaches for Extracellular Matrix Targeting in Disease Treatment (Nikolaos A. Afratis, Irit Sagi)....Pages 261-275
Back Matter ....Pages 277-278

Citation preview

Methods in Molecular Biology 1952

Davide Vigetti Achilleas D. Theocharis Editors

The Extracellular Matrix Methods and Protocols

METHODS

IN

MOLECULAR BIOLOGY

Series Editor John M. Walker School of Life and Medical Sciences University of Hertfordshire Hatfield, Hertfordshire, AL10 9AB, UK

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

The Extracellular Matrix Methods and Protocols

Edited by

Davide Vigetti Department of Medicine and Surgery, University of Insubria, Varese, Italy

Achilleas D. Theocharis Department of Chemistry, University of Patras, Patras, Greece

Editors Davide Vigetti Department of Medicine and Surgery University of Insubria Varese, Italy

Achilleas D. Theocharis Department of Chemistry University of Patras Patras, Greece

ISSN 1064-3745 ISSN 1940-6029 (electronic) Methods in Molecular Biology ISBN 978-1-4939-9132-7 ISBN 978-1-4939-9133-4 (eBook) https://doi.org/10.1007/978-1-4939-9133-4 Library of Congress Control Number: 2019932812 © Springer Science+Business Media, LLC, part of Springer Nature 2019 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, express 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 Press imprint is published by the registered company Springer Science+Business Media, LLC, part of Springer Nature. The registered company address is: 233 Spring Street, New York, NY 10013, U.S.A.

Preface Extracellular matrix (ECM) is a three-dimensional noncellular network where cells reside. Apart from structural support, ECM controls cell phenotype and functions in tissues and organs. ECM components are composed of fibrillar proteins including collagens, fibronectin, laminins and elastin, proteoglycans, hyaluronan, and several glycoproteins including matricellular proteins. All these ECM molecules interact with each other as well as with certain cell surface receptors creating complex but well-organized networks. Specifically, the elements of the ECMs can influence tissue homeostasis, cell proliferation, migration, survival, differentiation, angiogenesis, immune function, and autophagy. ECM is subjected in continuous remodeling in physiological and pathologic circumstances. ECM-degrading enzymes including matrix metalloproteinases and hyaluronidases are responsible for ECM remodeling, whereas resident cells synthesize and deposit newly formed ECM material to reconstitute the physiologic tissue architecture. ECM remodeling is regulated by cell-matrix and cell-cell communication which are mediated by direct and indirect interactions. Recently, exosomes have been demonstrated to be integral components of ECM that are involved in matrix remodeling and cell-matrix and cell-cell communication. Abnormal remodeling of ECM often occurs and is associated with disease development and progression. Researchers have focus on ECM molecules to develop new therapeutic modalities, drug delivery approaches, and biomaterials. Nowadays, ECM is considered as a regulatory element of tissues and organs and not only as a niche for cells. It controls cell fate and functions, and the research on matrix biology becomes equally important with that of cell biology. The aim of this volume of the Methods in Molecular Biology series is to provide readers with established experimental protocols useful for the isolation, characterization, and detection of ECM molecules as well as methods to study the activity and role of ECM components on various biological functions, the formation of exosomes and tissue surrogates. This volume is aimed at basic and clinical researchers including biochemists, cell and molecular biologists, pathologists, oncologists, and physicians. The detailed protocols and applications presented within this book series can be exploited by Ph.D. students and senior researchers to carry out research in the field of extracellular matrix biology. This book contains 21 chapters that display a variety of protocols ranging from biochemical and cell and molecular biology assays to complex tissue imaging techniques and in vivo models to elucidate the role of extracellular matrix. Chapter 1 provides a short introduction in ECM structure and its regulatory role on cell functions and behavior. It emphasizes on the complex interplay between ECM components and cells that controls tissue homeostasis and progression of diseases. The gene expression of ECM molecules directly on fixed tissues is described in Chap. 2. This easy to perform and convenient method allows the recognition of cells expressing a certain component and can be joined with immunostaining in the same tissue samples to demonstrate the biosynthesis of ECM in situ. Protocols on modern methods including transmission electron microscope, scanning electron microscope, and atomic force microscope to visualize the supramolecular assembly of collagen in tissues are provided in Chap. 3. Chapter 4 provides protocols for studying the nature, the posttranslational modifications, and the kinetic of secretion of collagen in vitro by using steady-state and pulse-chase labeling with 3H-proline. Protocols for the isolation and characterization of

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marine invertebrate-derived glycosaminoglycans as well as for the analysis of proteoglycan biosynthesis and secretion in vitro are presented in Chaps. 5 and 6. In Chap. 7, the methodology for the analysis of precursor UDP-sugars for the biosynthesis of various cellular glycans and glycoconjugates is described. It involves isolation from cultured cells and tissues utilizing ion-pair solid-phase extraction with graphitized carbon cartridges and subsequent analysis using anion-exchange high-performance liquid chromatography. Chapters 8 and 9 present methods for the determination of hyaluronan molecular mass utilizing simple electrophoresis and visualization of pericellular hyaluronan by a quick semiquantitative particle exclusion assay and validation by fluorescence detection. The methodology for the determination of cell surface-associated hyaluronan by flow cytometry using hyaluronan-binding proteins conjugated to different fluorochromes is described in Chap. 10. In Chap. 11, a protocol for isolation of primary cultures of adult rat small intestine myenteric ganglia and subsequent visualization of hyaluronan is provided. A simple and sensitive approach for analysis of hyaluronidases present in biological fluids employing hyaluronan zymography and microtiter plate assay is given in Chap. 12. Chapter 13 describes a protocol for studying the effect of glycosaminoglycans on voltage-dependent ion channel gating. Comprehensive methodology for monitoring matrix-induced autophagy utilizing morphological and microscopic characterization, biochemical analysis, and signaling pathway investigation as well as for detecting mitophagy via monitoring changes in mitochondrial DNA and permeability is provided in Chap. 14. In Chaps. 15 and 16, experimental approaches for determining gelatinolytic activity in tissues by in situ zymography and real-time gelatin zymography are described. Chapters 17 and 18 present methodologies for evaluation of angiogenesis by using in vitro and in vivo models. Chapter 19 includes a protocol for isolation and characterization of exosomes released in the cell culture medium. The methodology for the preparation and characterization of rich in extracellular matrix tissue surrogates from various cell populations using the principles of macromolecular crowding is described in Chap. 20. In addition, Chap. 21 discusses strategies to target ECM components for disease treatment. It proposes the creation of a pioneer ECM multidisciplinary integrated platform in order to decipher ECM homeostasis that can lead to the development of therapeutic new agents, protein-based inhibitors, diagnostics, medical devices, artificial tissues, and drug delivery strategies. We hope that this volume will be useful and enjoyable for readers and it will foster the advancement of research on extracellular matrix biology. We are very appreciative of all contributions provided by eminent scientists in the field. We are also grateful to Prof. John M. Walker, editor-in-chief of Methods in Molecular Biology series, for the continuous interest and support throughout this period that helped us to accomplish the editing of this volume. Varese, Italy Patras, Greece

Davide Vigetti Achilleas D. Theocharis

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

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1 The Complex Interplay Between Extracellular Matrix and Cells in Tissues . . . . . Dimitra Manou, Ilaria Caon, Panagiotis Bouris, Irene-Eva Triantaphyllidou, Cristina Giaroni, Alberto Passi, Nikos K. Karamanos, Davide Vigetti, and Achilleas D. Theocharis 2 Method for Studying ECM Expression: In Situ RT-PCR . . . . . . . . . . . . . . . . . . . . ` , Cristiana Marcozzi, Barbara Bartolini, Elena Carava Marcella Reguzzoni, Paola Moretto, Ilaria Caon, Evgenia Karousou, Alberto Passi, and Manuela Viola 3 Visualizing the Supramolecular Assembly of Collagen . . . . . . . . . . . . . . . . . . . . . . . Mario Raspanti, Marcella Reguzzoni, Petra Rita Basso, Marina Protasoni, and De´sire´e Martini 4 Steady-State and Pulse-Chase Analyses of Fibrillar Collagen . . . . . . . . . . . . . . . . . . Antonella Forlino, Francesca Tonelli, and Roberta Besio 5 Methods for Isolation and Characterization of Sulfated Glycosaminoglycans from Marine Invertebrates. . . . . . . . . . . . . . . . . . . . . . . . . . . . . Mariana P. Stelling, Ananda A. de Bento, Philippe Caloba, ˜o Eduardo Vilanova, and Mauro S. G. Pava 6 Analysis of Proteoglycan Synthesis and Secretion in Cell Culture Systems . . . . . . Chiara Paganini, Rossella Costantini, and Antonio Rossi 7 Analysis of UDP-Sugars from Cultured Cells and Small Tissue Samples. . . . . . . . Sanna Oikari and Markku I. Tammi 8 Methods for Hyaluronan Molecular Mass Determination by Agarose Gel Electrophoresis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Mary K. Cowman 9 Hyaluronan Pericellular Matrix: Particle Exclusion Assay. . . . . . . . . . . . . . . . . . . . . Melanie A. Simpson 10 Determination of Cell-Surface Hyaluronan Through Flow Cytometry. . . . . . . . . Daiana L. Vitale, Fiorella M. Spinelli, and Laura Alaniz 11 Method for Detecting Hyaluronan in Isolated Myenteric Plexus Ganglia of Adult Rat Small Intestine . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Michela Bistoletti, Paola Moretto, and Cristina Giaroni 12 Analyzing Hyaluronidases in Biological Fluids . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Christos Velesiotis, Stella Vasileiou, and Demitrios H. Vynios 13 Method for Studying the Physical Effect of Extracellular Matrix on Voltage-Dependent Ion Channel Gating . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Eleonora Solari and Andrea Moriondo 14 Methods for Monitoring Matrix-Induced Autophagy . . . . . . . . . . . . . . . . . . . . . . . Carolyn Chen, Aastha Kapoor, and Renato V. Iozzo

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71 81

91 103 111

117 127

143 157

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Method for Determining Gelatinolytic Activity in Tissue: In Situ Gelatin Zymography . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Elin Hadler-Olsen and Jan-Olof Winberg Method for Determining Gelatinolytic Activity in Tissue Extracts: Real-Time Gelatin Zymography . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Elin Hadler-Olsen and Jan-Olof Winberg In Vitro Spheroid Sprouting Assay of Angiogenesis . . . . . . . . . . . . . . . . . . . . . . . . . Fatema Tuz Zahra, Efrossini Choleva, Md Sanaullah Sajib, Evangelia Papadimitriou, and Constantinos M. Mikelis Matrigel Plug Assay for In Vivo Evaluation of Angiogenesis. . . . . . . . . . . . . . . . . . Pinelopi Kastana, Fatema Tuz Zahra, Despoina Ntenekou, Stamatiki Katraki-Pavlou, Dimitris Beis, Michail S. Lionakis, Constantinos M. Mikelis, and Evangelia Papadimitriou Exosomes from Cell Culture-Conditioned Medium: Isolation by Ultracentrifugation and Characterization. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Anurag Purushothaman Preparation and Characterization of Tissue Surrogates Rich in Extracellular Matrix Using the Principles of Macromolecular Crowding . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Adrian Djalali-Cuevas, Sergio Garnica-Galvez, Andrea Rampin, Diana Gaspar, Ioannis Skoufos, Athina Tzora, Nikitas Prassinos, Nikolaos Diakakis, and Dimitrios I. Zeugolis Novel Approaches for Extracellular Matrix Targeting in Disease Treatment . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Nikolaos A. Afratis and Irit Sagi

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

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Contributors NIKOLAOS A. AFRATIS  Department of Biological Regulation, Weizmann Institute of Science, Rehovot, Israel LAURA ALANIZ  Laboratorio de Microambiente Tumoral, Universidad Nacional del Noroeste de la Pcia. de Bs. As, Centro de Investigaciones y Transferencia del Noroeste de la Provincia de Buenos Aires (CITNOBA UNNOBA-CONICET), Junı´n, Buenos Aires, Argentina BARBARA BARTOLINI  Department of Medicine and Surgery, University of Insubria, Varese, Italy PETRA RITA BASSO  Department of Medicine and Surgery, University of Insubria, Varese, Italy DIMITRIS BEIS  Biomedical Research Foundation Academy of Athens, Athens, Greece ROBERTA BESIO  Biochemistry Unit, Department of Molecular Medicine, University of Pavia, Pavia, Italy MICHELA BISTOLETTI  Department of Medicine and Surgery, University of Insubria, Varese, Italy PANAGIOTIS BOURIS  Laboratory of Biochemistry, Department of Chemistry, University of Patras, Patras, Greece PHILIPPE CALOBA  Laboratorio de Bioquı´mica e Biologia Celular de Glicoconjugados, Programa de Glicobiologia, Instituto de Bioquı´mica Me´dica Leopoldo de Meis and Hospital Universita´rio Clementino Fraga Filho, Universidade Federal do Rio de Janeiro, Rio de Janeiro, Brazil; Instituto Federal de Educac¸a˜o, Cieˆncia e Tecnologia do Rio de Janeiro, Rio de Janeiro, Brazil ILARIA CAON  Department of Medicine and Surgery, University of Insubria, Varese, Italy ELENA CARAVA`  Department of Medicine and Surgery, University of Insubria, Varese, Italy CAROLYN CHEN  Department of Pathology, Anatomy and Cell Biology and the Cancer Cell Biology and Signaling Program, Sidney Kimmel Medical College at Thomas Jefferson University, Philadelphia, PA, USA EFROSSINI CHOLEVA  Laboratory of Molecular Pharmacology, Department of Pharmacy, School of Health Sciences, University of Patras, Patras, Greece ROSSELLA COSTANTINI  Department of Molecular Medicine, Biochemistry Unit, University of Pavia, Pavia, Italy MARY K. COWMAN  New York University, Tandon School of Engineering, New York, NY, USA ANANDA A. DE BENTO  Laboratorio de Bioquı´mica e Biologia Celular de Glicoconjugados, Programa de Glicobiologia, Instituto de Bioquı´mica Me´dica Leopoldo de Meis and Hospital Universita´rio Clementino Fraga Filho, Universidade Federal do Rio de Janeiro, Rio de Janeiro, Brazil; Instituto Federal de Educac¸a˜o, Cieˆncia e Tecnologia do Rio de Janeiro, Rio de Janeiro, Brazil NIKOLAOS DIAKAKIS  School of Veterinary Medicine, Aristotle University of Thessaloniki, Thessaloniki, Greece ADRIAN DJALALI-CUEVAS  Bioprocessing and Biotechnology Laboratory, Department of Agricultural Technology, Technological Education Institute (TEI) of Epirus, Arta, Greece; School of Veterinary Medicine, Aristotle University of Thessaloniki, Thessaloniki, Greece;

ix

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Contributors

Regenerative, Modular and Developmental Engineering Laboratory (REMODEL), National University of Ireland Galway (NUI Galway), Galway, Ireland ANTONELLA FORLINO  Biochemistry Unit, Department of Molecular Medicine, University of Pavia, Pavia, Italy SERGIO GARNICA-GALVEZ  Bioprocessing and Biotechnology Laboratory, Department of Agricultural Technology, Technological Education Institute (TEI) of Epirus, Arta, Greece; School of Veterinary Medicine, Aristotle University of Thessaloniki, Thessaloniki, Greece; Regenerative, Modular and Developmental Engineering Laboratory (REMODEL), National University of Ireland Galway (NUI Galway), Galway, Ireland DIANA GASPAR  Regenerative, Modular and Developmental Engineering Laboratory (REMODEL), National University of Ireland Galway (NUI Galway), Galway, Ireland; ´ RAM), Science Foundation Ireland (SFI) Centre for Research in Medical Devices (CU National University of Ireland Galway (NUI Galway), Galway, Ireland CRISTINA GIARONI  Department of Medicine and Surgery, University of Insubria, Varese, Italy ELIN HADLER-OLSEN  Faculty of Health Sciences, Department of Medical Biology, UiT—The Arctic University of Norway, Tromsø, Norway; Faculty of Health Sciences, Department of Clinical Dentistry, UiT—The Arctic University of Norway, Tromsø, Norway; Department of Clinical Pathology, University Hospital of North Norway, Tromsø, Norway RENATO V. IOZZO  Department of Pathology, Anatomy and Cell Biology and the Cancer Cell Biology and Signaling Program, Sidney Kimmel Medical College at Thomas Jefferson University, Philadelphia, PA, USA AASTHA KAPOOR  Department of Pathology, Anatomy and Cell Biology and the Cancer Cell Biology and Signaling Program, Sidney Kimmel Medical College at Thomas Jefferson University, Philadelphia, PA, USA NIKOS K. KARAMANOS  Laboratory of Biochemistry, Department of Chemistry, University of Patras, Patras, Greece EVGENIA KAROUSOU  Department of Medicine and Surgery, University of Insubria, Varese, Italy PINELOPI KASTANA  Laboratory of Molecular Pharmacology, Department of Pharmacy, School of Health Sciences, University of Patras, Patras, Greece STAMATIKI KATRAKI-PAVLOU  Laboratory of Molecular Pharmacology, Department of Pharmacy, School of Health Sciences, University of Patras, Patras, Greece; Biomedical Research Foundation Academy of Athens, Athens, Greece MICHAIL S. LIONAKIS  Fungal Pathogenesis Section, Laboratory of Clinical Immunology and Microbiology (LCIM), National Institute of Allergy & Infectious Diseases (NIAID), National Institutes of Health (NIH), Bethesda, MD, USA DIMITRA MANOU  Laboratory of Biochemistry, Department of Chemistry, University of Patras, Patras, Greece CRISTIANA MARCOZZI  Department of Medicine and Surgery, University of Insubria, Varese, Italy ´ DESIRE´E MARTINI  Department of Biomedical and Neuromotor Sciences, University of Bologna, Bologna, Italy CONSTANTINOS M. MIKELIS  Department of Pharmaceutical Sciences, School of Pharmacy, Texas Tech University Health Sciences Center, Amarillo, TX, USA PAOLA MORETTO  Department of Medicine and Surgery, University of Insubria, Varese, Italy ANDREA MORIONDO  Department of Medicine and Surgery, University of Insubria, Varese, Italy

Contributors

xi

DESPOINA NTENEKOU  Laboratory of Molecular Pharmacology, Department of Pharmacy, School of Health Sciences, University of Patras, Patras, Greece SANNA OIKARI  Institute of Biomedicine, University of Eastern Finland, Kuopio, Finland; Institute of Dentistry, University of Eastern Finland, Kuopio, Finland CHIARA PAGANINI  Department of Molecular Medicine, Biochemistry Unit, University of Pavia, Pavia, Italy EVANGELIA PAPADIMITRIOU  Laboratory of Molecular Pharmacology, Department of Pharmacy, School of Health Sciences, University of Patras, Patras, Greece ALBERTO PASSI  Department of Medicine and Surgery, University of Insubria, Varese, Italy MAURO S. G. PAVA˜O  Laboratorio de Bioquı´mica e Biologia Celular de Glicoconjugados, Programa de Glicobiologia, Instituto de Bioquı´mica Me´dica Leopoldo de Meis and Hospital Universita´rio Clementino Fraga Filho, Universidade Federal do Rio de Janeiro, Rio de Janeiro, Brazil NIKITAS PRASSINOS  School of Veterinary Medicine, Aristotle University of Thessaloniki, Thessaloniki, Greece MARINA PROTASONI  Department of Medicine and Surgery, University of Insubria, Varese, Italy ANURAG PURUSHOTHAMAN  Department of Lymphoma and Myeloma, The University of Texas MD Anderson Cancer Center, Houston, TX, USA ANDREA RAMPIN  Bioprocessing and Biotechnology Laboratory, Department of Agricultural Technology, Technological Education Institute (TEI) of Epirus, Arta, Greece; School of Veterinary Medicine, Aristotle University of Thessaloniki, Thessaloniki, Greece; Regenerative, Modular and Developmental Engineering Laboratory (REMODEL), National University of Ireland Galway (NUI Galway), Galway, Ireland MARIO RASPANTI  Department of Medicine and Surgery, University of Insubria, Varese, Italy MARCELLA REGUZZONI  Department of Medicine and Surgery, University of Insubria, Varese, Italy ANTONIO ROSSI  Department of Molecular Medicine, Biochemistry Unit, University of Pavia, Pavia, Italy IRIT SAGI  Department of Biological Regulation, Weizmann Institute of Science, Rehovot, Israel MD SANAULLAH SAJIB  Department of Pharmaceutical Sciences, School of Pharmacy, Texas Tech University Health Sciences Center, Amarillo, TX, USA MELANIE A. SIMPSON  Molecular and Structural Biochemistry, North Carolina State University, Raleigh, NC, USA IOANNIS SKOUFOS  Bioprocessing and Biotechnology Laboratory, Department of Agricultural Technology, Technological Education Institute (TEI) of Epirus, Arta, Greece ELEONORA SOLARI  Department of Medicine and Surgery, University of Insubria, Varese, Italy FIORELLA M. SPINELLI  Laboratorio de Microambiente Tumoral, Universidad Nacional del Noroeste de la Pcia. de Bs. As, Centro de Investigaciones y Transferencia del Noroeste de la Provincia de Buenos Aires (CITNOBA UNNOBA-CONICET), Junı´n, Buenos Aires, Argentina MARIANA P. STELLING  Laboratorio de Bioquı´mica e Biologia Celular de Glicoconjugados, Programa de Glicobiologia, Instituto de Bioquı´mica Me´dica Leopoldo de Meis and Hospital Universita´rio Clementino Fraga Filho, Universidade Federal do Rio de Janeiro, Rio de

xii

Contributors

Janeiro, Brazil; Instituto Federal de Educac¸a˜o, Cieˆncia e Tecnologia do Rio de Janeiro, Rio de Janeiro, Brazil MARKKU I. TAMMI  Institute of Biomedicine, University of Eastern Finland, Kuopio, Finland ACHILLEAS D. THEOCHARIS  Department of Chemistry, University of Patras, Patras, Greece FRANCESCA TONELLI  Biochemistry Unit, Department of Molecular Medicine, University of Pavia, Pavia, Italy IRENE-EVA TRIANTAPHYLLIDOU  Laboratory of Biochemistry, Department of Chemistry, University of Patras, Patras, Greece FATEMA TUZ ZAHRA  Department of Pharmaceutical Sciences, School of Pharmacy, Texas Tech University Health Sciences Center, Amarillo, TX, USA ATHINA TZORA  Bioprocessing and Biotechnology Laboratory, Department of Agricultural Technology, Technological Education Institute (TEI) of Epirus, Arta, Greece STELLA VASILEIOU  Biochemistry, Biochemical Analysis and Matrix Pathobiochemistry Research Group, Department of Chemistry, University of Patras, Patras, Greece CHRISTOS VELESIOTIS  Biochemistry, Biochemical Analysis and Matrix Pathobiochemistry Research Group, Department of Chemistry, University of Patras, Patras, Greece DAVIDE VIGETTI  Department of Medicine and Surgery, University of Insubria, Varese, Italy EDUARDO VILANOVA  Laboratorio de Bioquı´mica e Biologia Celular de Glicoconjugados, Programa de Glicobiologia, Instituto de Bioquı´mica Me´dica Leopoldo de Meis and Hospital Universita´rio Clementino Fraga Filho, Universidade Federal do Rio de Janeiro, Rio de Janeiro, Brazil MANUELA VIOLA  Department of Medicine and Surgery, University of Insubria, Varese, Italy DAIANA L. VITALE  Laboratorio de Microambiente Tumoral, Universidad Nacional del Noroeste de la Pcia. de Bs. As, Centro de Investigaciones y Transferencia del Noroeste de la Provincia de Buenos Aires (CITNOBA UNNOBA-CONICET), Junı´n, Buenos Aires, Argentina DEMITRIOS H. VYNIOS  Biochemistry, Biochemical Analysis and Matrix Pathobiochemistry Research Group, Department of Chemistry, University of Patras, Patras, Greece JAN-OLOF WINBERG  Faculty of Health Sciences, Department of Medical Biology, UiT—The Arctic University of Norway, Tromsø, Norway DIMITRIOS I. ZEUGOLIS  Regenerative, Modular and Developmental Engineering Laboratory (REMODEL), National University of Ireland Galway (NUI Galway), Galway, Ireland; Science Foundation Ireland (SFI) Centre for Research in Medical Devices ´ RAM), National University of Ireland Galway (NUI Galway), Galway, Ireland (CU

Chapter 1 The Complex Interplay Between Extracellular Matrix and Cells in Tissues Dimitra Manou, Ilaria Caon, Panagiotis Bouris, Irene-Eva Triantaphyllidou, Cristina Giaroni, Alberto Passi, Nikos K. Karamanos, Davide Vigetti, and Achilleas D. Theocharis Abstract Extracellular matrix (ECM) maintains the structural integrity of tissues and regulates cell and tissue functions. ECM is comprised of fibrillar proteins, proteoglycans (PGs), glycosaminoglycans, and glycoproteins, creating a heterogeneous but well-orchestrated network. This network communicates with resident cells via cell-surface receptors. In particular, integrins, CD44, discoidin domain receptors, and cell-surface PGs and additionally voltage-gated ion channels can interact with ECM components, regulating signaling cascades as well as cytoskeleton configuration. The interplay of ECM with recipient cells is enriched by the extracellular vesicles, as they accommodate ECM, signaling, and cytoskeleton molecules in their cargo. Along with the numerous biological properties that ECM can modify, autophagy and angiogenesis, which are critical for tissue homeostasis, are included. Throughout development and disease onset and progression, ECM endures rearrangement to fulfill cellular requirements. The main responsible molecules for tissue remodeling are ECM-degrading enzymes including matrix metalloproteinases, plasminogen activators, cathepsins, and hyaluronidases, which can modify the ECM structure and function in a dynamic mode. A brief summary of the complex interplay between ECM macromolecules and cells in tissues and the contribution of ECM in tissue homeostasis and diseases is given. Key words Extracellular matrix, Proteoglycans, Hyaluronan, Matrix metalloproteinases, Hyaluronidases, Syndecans, Integrins, Angiogenesis, Autophagy

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Introduction Tissues are constituted of various cells such as fibroblasts, epithelial and endothelial cells, pericytes and cells of the immune system, as well as of noncellular networks, the extracellular matrices (ECMs), within cells are embedded [1]. ECMs operate not only as a physical structural barrier for the cells, but also as a mean of communication, as they orchestrate crucial cellular functions through cell-cell and cell-matrix interactions [2]. Specifically, the elements of the ECMs can influence cellular proliferation, migration, survival,

Davide Vigetti and Achilleas D. Theocharis (eds.), The Extracellular Matrix: Methods and Protocols, Methods in Molecular Biology, vol. 1952, https://doi.org/10.1007/978-1-4939-9133-4_1, © Springer Science+Business Media, LLC, part of Springer Nature 2019

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differentiation, as well as tissue homeostasis, morphogenesis, angiogenesis, immune function, and autophagy [3–7]. The core macromolecules of ECMs are fibrillar proteins including collagens, fibronectin, laminins and elastin, proteoglycans (PGs), glycosaminoglycans (GAGs), and several glycoproteins including matricellular proteins, which altogether create these three-dimensional, complex but well-organized networks. Additionally, growth factors (GFs), cytokines, and matrix-remodeling enzymes coexist at this rich microenvironment [8] (Fig. 1). The components of the ECMs are synthesized and secreted by all cells to attribute cellular adaptability and support, whereas macromolecules can sequester the ECM mediators creating a directly mobilized source of signaling molecules such as growth factors and cytokines [5]. The composition of ECMs differs between tissues, development stages, and pathophysiological conditions. The content and structural features of matrix components segregate ECMs into interstitial and pericellular matrices, with the most representative example of the latter one to be the basement membrane [8]. The main molecules identified at the basement membrane are collagen IV, laminins, nidogens, and heparan sulfate PGs perlecan and agrin [9]. Interstitial matrices are dominated by fibrillar collagens, PGs, fibronectin, and matricellular proteins [8]. PGs are key players of the ECMs as they orchestrate numerous cellular functions. Different number and type of GAGs are covalently linked to the protein core of PGs. GAGs are heterogeneous negatively charged linear chains of repeating disaccharides that are constituted mostly by a hexosamine (N-acetyl-D-galactosamine or N-acetyl-D-glucosamine) and a hexuronic acid (D-glucuronic or Liduronic acid) or by galactose. GAGs are grouped in six categories: chondroitin sulfate (CS), dermatan sulfate (DS), heparan sulfate (HS), heparin (Hep), keratan sulfate (KS), and hyaluronan (HA). Except HA, all other GAGs can be sulfated at various positions of uronic acids, hexosamines, or galactose (KS) providing disaccharides and overall PGs with remarkable structural and functional diversity. According to the latest nomenclature, PGs are categorized based on their cellular and subcellular location, the gene and protein homology, and the existence of particular protein motifs within their protein core. So, PGs are classified as intracellular, with the only insofar characterized molecule to be serglycin, cell surface, pericellular, and extracellular. Cell-surface PGs depending on the manner that are attached to the cellular membrane are divided into transmembrane, including syndecans, CSPG4, betaglycan and phosphacan, and glycosylphosphatidylinositol (GPI) anchored represented by glypicans. Concerning pericellular PGs, perlecan, agrin, and collagen XVIII and XV are included. The last category of extracellular PGs contains the subfamily of hyalectans/lecticans with main representatives aggrecan and versican, the subfamily of

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Fig. 1 The heterogeneous ECM network performs orchestrated events to coordinate several crucial cell activities through signaling cascades. (1) Extracellular vesicles (EVs) are intrinsic constitutes of the ECM participating in the communication between cells. Their cargo consists of nucleic acids such as mRNAs, miRNAs, and noncoding RNAs, lipids, and proteins including adhesion, signaling, and cytoskeleton molecules. Their surface is rich in cell-surface receptors such as integrins, CD44, and HSPGs, as well as membraneassociated or membrane-bound proteases, thus facilitating their interplay with the ECM. (2) Endostatin and endorepellin, which are derived upon the proteolysis of the C-terminus of collagen XVIII and perlecan, respectively, regulate autophagy and angiogenesis via interaction with VEGFR and integrins. (3) Synthesis of HA is regulated by HA synthases (HAS) and its degradation by hyaluronidases. The fate of HA after its production is dual, as it can either be released in the ECM or remain closely attached to the cell membrane via its interaction with HAS and/or CD44. (4) Extracellular receptors intervene in the interplay between ECM elements and intracellular signals. Specifically, CD44 functions as co-receptor for GFs and cytokines, while DDRs are mostly associated with collagens. The role of integrins is critical as they interact with laminin, collagens, fibronectin, and HSPGs, as well as with actin cytoskeleton. (5) Hyalectans such as versican isoforms (e.g., V0, V1, V2, and V3) and aggrecan interact with HA, creating large aggregates which hold space integrity and water-retaining properties. (6) Collagens are abundant in the ECM providing structural support to the cells, due to their fine organized meshes which are supported by small leucine-rich PGs (SLRPs). (7) Voltage-gated ion channels (VGICs) additionally to their function of selective administration of ions can interact with laminin, GFs, and integrins, but also with actin cytoskeleton. (8) Matricellular proteins interact with GFs and cell-surface receptors such as integrins and HSPGs. (9) Upon the action of enzymes including MMPs, ectodomains of shed syndecans are liberated and diffused in the ECM resulting in the creation of chemotactic gradients. (10) ECM reorganization occurs due to the action of proteases, such as MMPs, ADAMs, ADAMTS, plasminogen activators, and plasmin, while acidic conditions favor the activity of secreted cathepsins. (11) Syndecans and glypicans act as co-receptors presenting growth factors (GFs) to their cognate receptors

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small leucine-rich proteoglycans (SLRPs) with decorin, biglycan, and lumican as prototype members and SPOCK [10] (Fig. 1). The 28 members of collagens consist of the most abundant and complex structures in mammalians and, in particular, represent the one-third of overall proteins in humans. Depending on the domain homology and function, collagens are divided into seven groups: fibrillar collagens, network-forming and beaded filament-forming collagens, anchoring fibrils, fibril-associated collagens with interrupted triple helices (FACITs), membrane-associated collagens with interrupted triple helices (MACITs), and multiple triplehelix domains and interruptions (MULTIPLEXIN)/endostatinproducing collagens [8, 11]. Of special interest are the additional elements on the collagen fibrils which can be calcium compounds involved in biomineralization of collagens and small PGs such as SLRPs that participate in collagen fibrillogenesis [12] (Fig. 1). Fibronectin is a glycoprotein that is expressed ubiquitously into the ECM and can be found either as dimer or as fibrils possessing pivotal roles in matrix assembly. Other important proteins of the ECM are laminins and matricellular proteins which are involved in crucial functions for tissue homeostasis and wound healing [8] (Fig. 1).

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ECM-Cell Communication Cells in order to orchestrate their multiple functions rely on extracellular receptors, such as integrins, discoidin domain receptors (DDRs), cell-surface PGs, and CD44. Integrins are the major receptors responsible for cell adhesion due to their ability to bridge the actin cytoskeleton with ECM components and enhance signaling cascades, mainly through focal adhesion kinase (FAK) and Src tyrosine kinases triggering outside-in signaling. Integrins are constituted from two heterodimeric subunits, α and β, with the β subunit to be responsible for inside-out signaling through the binding to intracellular activators, such as talin [8, 13] (Fig. 1). This mechanism regulates the affinity of integrins with ECM elements and thus cellular migration and adhesion [14]. Integrins are expressed by tumor and stromal cells and they are important for their communication with the tumor microenvironment [15, 16]. Especially, cancer-associated fibroblasts (CAFs), which are responsible for the majority of ECM alterations in the tumor microenvironment, are depended on integrins to mediate their functions, mostly through the activation of transforming growth factor β (TGF-β) and other growth factor signaling cascades [13]. DDRs belong to receptor tyrosine kinases, and nevertheless hold distinguished characteristics such as their mechanism of activation and ability to bind ECM macromolecules, and especially collagens. DDRs exist in two closely related receptors, DDR-1

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and DDR-2, with the last to be expressed in mesenchymal cells, while the first in epithelial cells [17, 18]. DDRs regulate crucial cellular functions, such as proliferation, differentiation, and migration in physiological conditions and their deregulation contributes to diseases, such as fibrosis and cancer [18, 19] (Fig. 1). CD44 can act as receptor, part-time PG, and regulator of signal transduction. Due to its canonical Ig motif, CD44 operates as receptor for GFs, cytokines, matrix metalloproteinases (MMPs), and HA [20]. HS and/or CS, attached to CD44, bind vascular endothelial growth factor (VEGF) and hepatocyte growth factor (HGF), while its cytoplasmic domain is responsible for the binding of intracellular regulatory proteins [21, 22]. Variants of CD44 as well as the standard form contribute to tumorigenesis [23–25] as well as to tumor-initiate capacity of cancer stem cells (CSCs) [26, 27] (Fig. 1). Interestingly, syndecans, as transmembrane proteoglycans, can transmit intracellular signals. Syndecans comprise of four members and are ubiquitously expressed in cells [28]. Six members of glypicans are also widely expressed in tissues and together with syndecans are involved in cell-matrix interactions [29]. Syndecans and glypicans mainly carry HS chains in which a plethora of ECM components containing Hep/HS-binding motifs can bound. Cellsurface PGs including syndecans and CSPG4 cooperate with integrins for binding to numerous ECM components activating signaling pathways that regulate cell functions. Syndecans and glypicans are also characterized by their ability to function as co-receptors due to their interaction with several GFs and cytokines regulating their signaling [8, 13]. Upon the action of sheddases, soluble fragments of syndecans can be administrated in the extracellular space, with simultaneous release of their binding partners, thus promoting cell migration and adhesion [30] (Fig. 1). Recently extracellular vesicles (EVs) have been recognized as integral components of ECM that regulate cell-cell and cell-matrix interactions [31]. EVs are structures formed by a lipid bilayer, wherein plenty of nucleic acids, lipids, and proteins are stored. In particular, mRNAs, miRNAs, noncoding RNAs, as well as molecules responsible for antigen presentation, adhesion, signaling, and cytoskeleton configuration can be encapsulated (Fig. 1). Extracellular vesicles depending on their origin are divided into exosomes with a size variation from 30 to 150 nm and microvesicles which are about 1 μm, whereas a biologically distinct category is the apoptotic bodies with size between 1 and 5 μm. Exosomes are created via an endocytic pathway, while microvesicles are products of the cell surface [32–34]. EVs are implicated in the mineralization procedures of cartilage, bone, and dentin, in tissue repair and wound healing, as well as in tumor growth and angiogenesis. EVs can directly be involved in signal transduction via interaction with surface receptors or fusing with the plasma membrane of the targeted cells, but also through EV endocytosis and release of their

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cargo within endosomal/lysosomal compartments [31]. A critical interplay exists between ECM and EVs, as the former regulates the biogenesis, intrusion, and destination of EVs and the latter are responsible for ECM reorganization. This last feature is mediated by stored or surface-associated enzymes such as membrane-type MMPs (MT-MMPs) (e.g., MT1-MMP), a disintegrin and metalloproteinase (ADAMs) (e.g., ADAM-17), a disintegrin and metalloproteinase with thrombospondin motifs (ADAMTs), hyaluronidases, sialidases, and heparanase, which cleave a variety of ECM constituents and/or act as sheddases producing soluble biologically active protein fragments [35, 36]. For example, heparanase owns multiple roles in manipulating the behavior of EVs, as it can enhance their secretion and be present as an exosome cargo, but also as a surface-associated enzyme can degrade HS chains and thus facilitate the penetration of exosomes through the local microenvironment. HA and HSPGs/CSPGs attached to exosome membrane regulate several functions, due to interaction with cell-surface receptors, such as CD44 and integrins, respectively [31, 36] (Fig. 1). Voltage-gated ion channels (VGICs) are also implicated in ECM-cell interactions since they selectively administer Na+, K+, Ca+2, and Cl
through the cell membrane in response to mechanical stimuli. Afterwards, the mechanical force is converted into electrical and biochemical signals, regulating pathophysiological conditions related to skeletal muscles and the cardiovascular and the nervous system. VGICs interact with ECM components and receptors such as integrins in vascular smooth muscle, fibronectin, and nerve growth factor (NGF) in molluscan neurons and laminin in atrial myocytes. Cytoskeleton plays a dual role in VGIC function as actin disorganization in retinal bipolar cells results in activation of VG K+ channel, but inhibits the L-type Ca+2 channels in vascular smooth muscle and modifies the time course of activation of the Na+ channel in cardiac tissue [37]. Additionally, CS can induce the activation of VGICs, either due to their ability to sequester calcium, thus influencing the outer membrane potential, or due to the activation of intracellular pathways [38]. Intracellularly, signals through VGICs can spread via GTPases such as Rho, Rac, and Cdc42, whereas Ras is probably involved in the signal transduction through VG Ca+2 current by NGF and Src kinase in dorsal root ganglia neurons [37]. It has also been shown that the blockage of VG Ca+2 channels resulted in attenuation of metastatic cell behaviors in an in vivo breast cancer model [39] (Fig. 1).

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Hyaluronan, a Unique Matrix Component: Biosynthesis and Functions HA is a ubiquitous component of ECM of mammals and other vertebrate animals that, during evolution, needed a molecule able

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to fill space in tissues and, in addition, also able to modulate the immune system [40]. HA is composed by D-glucuronic acid (GlcUA) and N-acetyl-D-glucosamine (GlcNAc) linked via alternating β-(1 ! 4) and β-(1 ! 3) glycosidic bonds. Differently from other GAGs, no further chemical modifications are observable in HA. Intriguingly, this simple polysaccharide shows many important physiological and pathological functions [41–43]. One of the most interesting features of HA is the ability to modulate immune system [44]; this property depends on the polysaccharide molecular mass (i.e., HA polymer length) [45, 46], on the chemical modification of HA with the two heavy chains that are present on the serum proteoglycan inter-alpha inhibitor mediated by the tumor necrosis factor-stimulated gene 6 (TSG6) [47, 48], and on the capacity to bind to different surface receptors as CD44 and Toll-like receptors 2/4 (TLR2/4) [41, 49, 50]. In ECM, HA can interact with several HA-binding proteins and proteoglycans determining several tissue properties as hydration and stiffness as recently reported [51, 52]. In tissues the length of the HA chains depends on the fine balancing between the synthetic and the catabolic processes (due to hyaluronidases) as well as the presence of oxidants, for example reactive oxygen species or UV light, that are able to fragment the HA polymers [41] (Fig. 1). In vivo, the final effect of these processes is to generate HA molecules of different lengths leading to polymers of different sizes varying from 6 to 7  106 Dalton (high-molecular-weight HA) to 1  106 Dalton (low-molecular-weight HA) [46]. It is widely accepted that high-molecularmass HA has an anti-inflammatory effect whereas, on the contrary, low-molecular-mass HA has pro-inflammatory properties [44]. The synthesis of HA is a complex process that, at the end, will produce the polysaccharide that can be released in the ECM or, in turn, can remain associated to the cell membrane [53]. This latter point is important for the determination of the pericellular space or coat and can be achieved through the binding to HA surface receptors, or the nascent HA polymer can remain associated to the synthetic enzymes (named HA synthase (HAS) 1, 2, and 3) outside the plasma membrane [54]. In mammals, HA synthesis is carried out in cells by the three isoenzymes HAS1, 2, and 3 that are located on the plasma membrane (Fig. 1). Although the crystal structure of mammals HAS enzymes is not yet available, several studies clarify the topological organization of class I HAS proteins that consist of six to eight transmembrane helices, which form a large cytosolic loop that is critical for catalysis. Recently, important structural information can be obtained by comparing HAS and chitin synthase [55]. HAS are complex enzymes; in fact, a HAS protein should (1) bind two distinct UDP sugars, (2) transfer two different sugars into two different glycosidic linkages, (3) repeat sugar polymerization, and (4) HA transfer across the membrane. Moreover, such protein should be somehow able to control the

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length of the nascent polysaccharide [56]. In mammals the most important and highly expressed HAS isoenzyme is HAS2; it is also reported that HAS2 expression and activity are finely tuned (for a review see [57]). Interestingly, some pathological microorganisms acquired the capability to synthesize a HA capsule that is used to escape the host immune system [58]. Although HA is simply composed by the disaccharide GlcUA-GlcNAc, the synthesis of these two sugars is quite peculiar. UDP-GlcUA and UDP-GlcNAc are two sugar nucleotides that are the substrates for HA synthesis in all HAS [59]. UDP sugars are also the building blocks used for the synthesis of all the other GAGs that, differently from HA, are produced inside the Golgi apparatus. Several sugar transporters are located on the Golgi membrane to allow the entrance of such substrates in the Golgi lumen, and, as the affinity and efficiency of such transporters are usually high, it is important to highlight that the concentration of UDP sugars in the Golgi could be different from that in the cytoplasm [60]. UDP sugar availability in the cytoplasm is a critical factor for the regulation of HA synthesis [61–64]. Hyaluronidases (HYALs) are a family of enzymes that catalyze HA degradation (Fig. 1). There are different classes of HYALs based on the reaction mechanism and typically mammal HYALs are hydrolases whereas prokaryotic HYALs belong to lyase enzymes that introduce a double bond in the reaction product. This unsaturated disaccharide can be easily derivatized and used in HA quantification [65]. In mammals, there are five functional HYALs: HYAL1, HYAL2, HYAL3, HYAL4, and HYAL5 (also known as SPAM1 or PH-20). HYAL6 (also known as HYALP1) is a pseudogene [66]. HYAL1 and HYAL2 are the most important HYALs in the majority of tissues. HYAL2 is a GPI-anchored protein and is responsible for cleaving high-molecular-weight HA, which is mostly bound to the CD44 receptor. After the CD44-mediated endocytosis, the resulting HA fragments are hydrolyzed by HYAL1 [66]. Interestingly, HYALs have a very acidic optimal pH; recently, KIAA1199 (alternative names CEMIP and HYBID) has been identified as a new member of enzyme able to degrade HA at neutral pH [67]. Being ubiquitous and one of the most abundant components of the ECM, HA is involved in many pathophysiological processes in mammals. HA can have different functions, but, essentially, it can serve as (1) structural component by organizing the ECM via the binding with other proteoglycans as aggrecan (typically in cartilage) or versican (in the vasculature or in dermis) or other HA-binding proteins named hyaladherins [68]; (2) space-filling molecule as in synovial fluids where, thanks to the mechanical proprieties as viscosity, HA has a lubricant function allowing cartilage movements or in vitreous humor of the eye where HA forms a gel structure with other ECM components; and (3) signaling molecule interacting

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with several membrane receptors as CD44, RHAMM, LYVE-1, Layilin, and HARE [49]. These receptors are used for the physiological turnover and degradation of HA, but are also critical to trigger specific signals that modulate survival, proliferation, and migration (these effects are particularly studied in several types of malignancies) [69, 70] as well as differentiation/dedifferentiation (i.e., in the phenotypic switch from contractile to synthetic in vascular smooth muscle cells or in the switch from fibroblasts to myofibroblasts) [71] and angiogenesis (i.e., in lung epithelium or endothelial cells) [72] (Fig. 1). Another important factor is the molecular mass of HA that can have a dramatic impact on signaling. As described above low-molecular-mass HA (3000 kDa)). We usually use HA in a 0.15 M NaCl solution, with or without a little phosphate buffer, but it appears that deionized water is also fine. Mix 5 μL of HA sample (containing approximately 2.5 μg HA) with 10 μL water and 3 μL of 0.02% bromophenol blue (optional) and 2 M sucrose in TBE. If the HA solution is more dilute than 0.3–0.5 mg/mL, decrease the water accordingly. If the HA solution is much more concentrated, it may need to be diluted to 0.5 mg/mL at least one day in advance so that chains can fully disentangle. 2. HA standards: Mix 5 μL of the HA standard solution (containing 1 μg HA) with 10 μL water and 3 μL of 0.02% bromophenol blue (optional) and 2 M sucrose in TBE.

3.1.3 Electrophoresis

1. Remove the comb from the gel while it is still covered by the TBE buffer. 2. Make sure that the gel electrophoresis unit is leveled. Carefully, with both hands, pick up the gel from the casting apparatus and let excess buffer flow off. Place the gel in the center of the electrophoresis unit. 3. Fill the electrophoresis unit with ca. 245 mL 1 TBE buffer (should already be at room temperature). It should result in a ca. 4 mm thick layer of buffer above the gel (see Note 18). 4. Apply the standard and samples into empty wells. Use a 1–20 μL repeating pipette. Adjust the pipette to deliver 18 μL. Gently remix the sample, and then slowly pipette each sample to the bottom of the specified well so that it is layered under the buffer. Do not create turbulence that may mix the sample with the electrode buffer. Record the lane number of each sample. Standards are usually placed in the first or last well. When the gel is to be used for quantitative analysis of HA molecular mass distribution, each sample must be adjacent to an empty lane, for background subtraction of stain. 5. Place the cover on the apparatus and connect wires to the proper outlet of the power source. The negative electrode

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(black wire) must always be connected to the end of the apparatus that is closest to the top of the gel where the wells are located, and the positive electrode (red wire) must always be connected to the end that is closest to the bottom of the gel. 6. Electrophorese at 20 V constant voltage for 0.5 h, and then 40 V for 3.5 h. The bromophenol blue dye migrates almost to the end of the gel in this time. 7. Immediately after the current has been turned off, the gel must be removed from the gel apparatus and placed in the staining solution. Sample diffusion may occur if there is a delay between these two steps. 3.1.4 Staining and Destaining

1. Pour a little Stains-All dye into a rectangular glass dish to wet the bottom so the gel will not stick. 2. Gently and carefully transfer the gel from the electrophoresis unit to the rectangular glass dish (see Note 19). 3. Slowly pour ca. 500 mL of Stains-All solution to the stain dish, or enough to cover the gel (depends on the size of the glass dish). Remove all bubbles that may be trapped under the gel, and make sure that it is freely moving in the stain solution by rocking the dish back and forth a few times. 4. Cover the dish with plastic wrap. Wrap the entire dish (top and bottom, in two pieces) with aluminum foil to protect it from light. Put in a dark cabinet at room temperature overnight. 5. Destain the gel by removing all the Stains-All solution (use suction created with a lab vacuum source connected to a 1 L Erlenmeyer flask trap, and flexible silicone rubber tubing to immerse in the stain). Add about 500 mL of 10% ethanol to the dish. Tilt dish back and forth to ensure that the gel is not stuck to the dish. Re-cover with plastic wrap and foil and put in dark cabinet. It is best to change the destaining solution at least once. The destaining can be done overnight or preferably for approximately 1 day. 6. Use a photocopier transparency sheet (cut to fit inside the glass dish but larger than the gel) to slip under the gel and lift it out of the glass dish. Use both hands; the gel will try to escape. Let the destaining solution run off the gel a little. The bottom of the transparency can be laid briefly on a paper towel to dry it. 7. Place the gel and transparency sheet on a light box to fade the remaining unbound dye. The purple color of the background will fade in a few minutes. Do not leave the gel too long or the sample bands will also fade. If there is any apparent precipitated dye adhering to the gel surface, squirt a little 10% ethanol on it and rub gently with a gloved finger.

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8. The gel can preferably be scanned at this point, or can be returned to a fresh 10% ethanol solution for storage in the dark for several days. 9. Discard all waste material in accordance with current corporate or university policy. Stains-All solution and destaining solutions must be handled as hazardous waste. Charcoal filters may prove helpful in removing the dye from solutions. Gels containing HA samples from human tissues should be handled as biohazard material. 3.2 Agarose Gel Electrophoresis in TAE Buffer and Staining

3.2.1 TAE Gel Preparation

This is the procedure for a short (10 cm long) gel, with 0.5% agarose in TAE. The gel is cast in the casting accessory for the Bio-Rad Mini Sub Cell GT. The gel dimensions are 10 cm long and 6.2 cm wide. The agarose gel volume is 40 mL, so the gel thickness is ca. 6.5 mm. An eight-tooth well-forming comb is set at a height leaving 1 mm clearance from the bottom, creating wells of 1  5  5.5 mm. The gel will be run in a much larger horizontal electrophoresis unit, such as the LKB 2012 Maxiphor (or similar) with dimensions of ca. 20 cm long  15 cm wide. The reason for this is that use of the small electrophoresis unit results in buffer exhaustion during the run, and the pH falls significantly at the end of the gel (near the positive electrode), resulting in poor migration and poor staining of the fastest HA. 1. 0.5% Agarose solution: Weigh 0.2 g agarose into a 125 mL Erlenmeyer flask. Add 36 mL deionized water. Cover with Parafilm. Use a microwave oven to heat up the solution to help it dissolve. A total time of perhaps 90 s at full power, interrupted a few times to swirl the flask (whenever the agarose solution starts to boil), may be needed. Stop when the solution is clear and free of all particles, including any nearly clear gel-like bits of agarose. Transfer the flask to a 48  C water bath for 15 min. You will need to put on new Parafilm. Prewarm 4 mL 10 TAE buffer at 48  C for the same 15 min. Add the 4 mL prewarmed 10 TAE buffer into the agarose solution. Swirl to mix well. 2. Pouring the gel: Follow the procedure in Subheading 3.1.1, step 2, substituting TAE buffer for TBE buffer.

3.2.2 Sample Preparation 3.2.3 Electrophoresis

Follow the procedure described in Subheading 3.1.2.

1. Remove the comb from the gel while it is still covered by the TAE buffer. 2. Make sure that the large format gel electrophoresis unit is leveled. Carefully, with both hands, pick up the gel from the

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casting apparatus and let excess buffer flow off. Place the gel in the center of the large electrophoresis unit. 3. Fill the electrophoresis unit with ca. 1200 mL 1 TAE buffer (should already be at room temperature). It should result in a ca. 3 mm thick layer of buffer above the gel. This is very important. Too much or too little buffer above the gel will harm the separation. Made sure that the gel is not floating. If bubbles are seen under the gel tray, the tray is lifted up and tilted slowly to get rid of bubbles, and then replaced. 4. Apply samples into empty wells. Follow the procedure in Subheading 3.1.3, step 4. 5. Place the cover on the apparatus and connect wires to the proper outlet of the power source. Follow the procedure in Subheading 3.1.3, step 5. 6. Electrophorese at 20 V constant voltage for 0.5 h, and then 40 V for 3.5 h. The bromophenol blue dye migrates almost to the end of the gel in this time. 7. Immediately after the current has been turned off, the gel must be removed from the gel apparatus and placed in the staining solution. See Subheading 3.1.3, step 7. 3.2.4 Staining and Destaining

3.3 Densitometry and Analysis of Stained Agarose Gels 3.3.1 Densitometry

3.3.2 Data Analysis

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Follow the procedure described in Subheading 3.1.4.

1. Quantitative analysis of gels should be done with a densitometric scanner operating in transmission mode, such as ImageScanner or ImageScanner III from GE Healthcare, using LabScan software. 2. The gel can be imaged or scanned using a red filter to enhance the blue stain. An example of the scanned gel image for a 0.5% agarose gel in TAE buffer is shown in Fig. 1. 1. Detailed procedures and appropriate template spreadsheet files are available as Supplementary Data files 1–3 in reference [7].

Notes 1. 121 g Tris base is dissolved in 800 mL H2O in 1 L beaker. Add dry boric acid (approx. 60 g) slowly with stirring to pH 8.3. Transfer it to 1 L volumetric flask. Adjust volume to 1 L. 2. 7.44 g Na2EDTA dihydrate is dissolved in 80 mL H2O in a 150 mL beaker. Adjust pH to approx. 7 with 10 M NaOH. Transfer to 100 mL volumetric flask. Adjust volume to 100 mL.

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Fig. 1 Electrophoresis of HA on 0.5% agarose in TAE buffer, stained with StainsAll dye. The chemoenzymatically synthesized HA standards (Select-HA) appear as sharp bands, but HA isolated from tissue or bacterial sources appears highly polydisperse. The viscosity-average molecular mass of the two polydisperse samples is indicated. This figure has been adapted with permission from Cowman, M. K.; Chen, C. C.; Pandya, M.; Yuan, H.; Ramkishun, D.; LoBello, J.; Bhilocha, S.; Russell-Puleri, S.; Skendaj, E.; Mijovic, J.; Jing, W. Improved Agarose Gel Electrophoresis Method and Molecular Mass Calculation for High Molecular Mass Hyaluronan. Anal. Biochem. 2011, 417, 50–56. Copyright 2011, Elsevier Inc

3. Mix 10 mL 1 M Tris-borate, pH 8.3, and 0.5 mL 0.2 M Na2EDTA, pH 7, in a 100 mL graduated cylinder. Add deionized water to make the total volume 50 mL. Mix well using magnetic stir bar. Make this buffer solution only when needed for sample loading and tracking dye solutions. 4. Mix 100 mL 1 M Tris-borate, pH 8.3, with 5 mL 0.2 M Na2EDTA, pH 7, and 895 mL H2O. 5. 6.84 g Sucrose is dissolved in 5 mL 2 TBE, volume adjusted to 10 mL with H2O. 6. 0.002 g Bromophenol blue and 6.84 g sucrose are dissolved in 5 mL 2 TBE buffer, volume adjusted to 10 mL with H2O. 7. Use a 2 L graduated cylinder. Mix 1 L absolute (100%, 200 proof, but not denatured) ethanol with 1 L H2O. If you use 95% ethanol, adjust volumes accordingly.

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8. Weigh 0.05 g Stains-All and transfer to a 1 L volumetric flask. Add 50% ethanol solution and adjust the volume to 1 L. Add a stir bar and cover the flask with aluminum foil to prevent photodegradation and consequent fading of the dye. Stir for at least 1.5 h. We sometimes, but not normally, find it necessary to filter the Stains-All solution (in a dimly lit room) through Whatman #1 filter paper before use. This is necessary if the dye is heavily contaminated with photodegradation products, which are less soluble. 9. Use a 2 L graduated cylinder. Mix 200 mL absolute ethanol with 1800 mL DI water. If you use 95% ethanol, adjust volumes accordingly. 10. HA standards of known molecular mass and very low polydispersity produced chemoenzymatically according to published methods [10, 11] have been commercialized by Hyalose LLC. These are supplied in a dry form and must be dissolved in (preferably 0.2 μm prefiltered) deionized water. Follow the directions supplied. Add the water but do not mix in any way the first day. Let it dissolve in the refrigerator. The second day, gently pipette up and down to mix and wash down the sides of the tube. Store in the refrigerator. Remix with micropipette before use. If this product is unavailable, the published methods can be used to prepare standards. Alternatively, polydisperse HA can be fractionated by gel filtration or ion-exchange chromatography to obtain standards. These standards should ideally be characterized by light scattering for absolute determination of average molecular mass. As a less desirable solution, the correlation of mobility between HA of known molecular mass and DNA restriction fragments can be used [5], with the full understanding that these molecules have different inherent mobility for the same molecular mass, and the correlations are only useful when cross-calibrated. 11. Weigh into a 1 L beaker 48.4 g Tris base, 6.8 g sodium acetate trihydrate, and 3.3 g disodium EDTA dihydrate. Add about 750 mL H2O. Use a magnetic stir bar to stir until dissolved. Adjust pH to 7.9, while stirring, using dropwise addition of concentrated (glacial) acetic acid. Transfer to 1 L volumetric flask (rinse beaker with water and add to volumetric). Adjust volume to 1 L. Alternative procedure for 10 TAE buffer: Weigh into a 2 L beaker 36.3 g Tris base, 78.4 g Tris–HCl, 13.6 g sodium acetate trihydrate, and 6.6 g disodium EDTA dihydrate. Add about 1500 mL H2O. Use a magnetic stir bar to stir until dissolved. Check pH. It should be 7.9. If needed, adjust pH to 7.9 using HCl or NaOH. Transfer to 2 L volumetric flask (rinse beaker with water and add to volumetric). Adjust volume to 2 L. Store at 4  C. Discard after 1 month.

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12. Dilute 10 mL 10 TAE buffer with H2O to make the total volume 50 mL. Mix well using magnetic stir bar. Make this buffer solution only when needed for sample loading and tracking dye solutions. 13. Dilute 200 mL of 10 TAE with 1800 mL H2O to 2 L. This can be done in a 2 L graduated cylinder. Cover with plastic wrap or Parafilm. 14. 6.84 g Sucrose is dissolved in 5 mL 2 TAE, volume adjusted to 10 mL with H2O. 15. 0.002 g Bromophenol blue and 6.84 g sucrose dissolved in 5 mL 2 TAE buffer, volume adjusted to 10 mL with H2O. 16. There are many sources for agarose, but they can vary in purity and therefore vary in how well they work for this purpose. The agarose should have a low electroendosmosis number. This number reflects the fraction of anionic groups on the agarose. This is important for avoidance of band smearing due to the migration of mobile cations and the accompanying flow of water in the reverse direction to HA migration. It is also important for reduction of background staining in the gel matrix. We most commonly use Agarose NA from GE Healthcare. We have observed that Stains-All dye can be precipitated by some less pure agarose. 17. Some people put gels in the cold overnight or for longer times but it should be at room temperature for the run. Some people use the gels the same day they are poured. We prefer overnight incubation at room temperature. 18. The depth of the buffer above the agarose gel is very important. Too much or too little buffer above the gel will harm the separation. Make sure that the gel is not floating. If bubbles are seen under the gel tray, the tray is lifted up and tilted slowly to get rid of bubbles, and then replaced. 19. The gel tray is slowly taken out of the electrophoresis unit, tilting it slightly to let buffer run off, but being careful to stop the gel from sliding off. With a Kimwipe tissue, wipe the bottom and edges of the tray to avoid excess buffer contamination of the stain solution. Hold the gel tray at a 45-degree angle to slide the gel into the glass dish. If it sticks a little, use a gloved finger to gently push it off. References 1. Knudson W, Gundlach MW, Schmid TM, Conrad HE (1984) Selective hydrolysis of chondroitin sulfates by hyaluronidase. Biochemistry 23(2):368–375

2. Hampson IN, Gallagher JT (1984) Separation of radiolabelled glycosaminoglycan oligosaccharides by polyacrylamide-gel electrophoresis. Biochem J 221(3):697–705

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3. Cowman MK, Slahetka MF, Hittner DM, Kim J, Forino M, Gadelrab G (1984) Polyacrylamide-gel electrophoresis and Alcian Blue staining of sulphated glycosaminoglycan oligosaccharides. Biochem J 221(3):707–716 4. Min H, Cowman MK (1986) Combined Alcian blue and silver staining of glycosaminoglycans in polyacrylamide gels: application to electrophoretic analysis of molecular weight distribution. Anal Biochem 155(2):275–285 5. Lee HG, Cowman MK (1994) An agarose gel electrophoretic method for analysis of hyaluronan molecular weight distribution. Anal Biochem 219(2):278–287 6. Bhilocha S, Amin R, Pandya M, Yuan H, Tank M, LoBello J, Shytuhina A, Wang W, Wisniewski HG, de la Motte C, Cowman MK (2011) Agarose and polyacrylamide gel electrophoresis methods for molecular mass analysis of 5- to 500-kDa hyaluronan. Anal Biochem 417(1):41–49 7. Cowman MK, Chen CC, Pandya M, Yuan H, Ramkishun D, LoBello J, Bhilocha S, RussellPuleri S, Skendaj E, Mijovic J, Jing W (2011) Improved agarose gel electrophoresis method and molecular mass calculation for high molecular mass hyaluronan. Anal Biochem 417 (1):50–56 8. Ikegami-Kawai M, Takahashi T (2002) Microanalysis of hyaluronan oligosaccharides by polyacrylamide gel electrophoresis and its application to assay of hyaluronidase activity. Anal Biochem 311(2):157–165 9. Cowman MK (2017) Hyaluronan and hyaluronan fragments. Adv Carbohydr Chem Biochem 74:1–59 10. Jing W, DeAngelis PL (2004) Synchronized chemoenzymatic synthesis of monodisperse hyaluronan polymers. J Biol Chem 279 (40):42345–42349 11. Jing W, Haller FM, Almond A, DeAngelis PL (2006) Defined megadalton hyaluronan polymer standards. Anal Biochem 355(2):183–188 12. Armstrong SE, Bell DR (2002) Measurement of high-molecular-weight hyaluronan in solid tissue using agarose gel electrophoresis. Anal Biochem 308(2):255–264

13. Takeo S, Fujise M, Akiyama T, Habuchi H, Itano N, Matsuo T, Aigaki T, Kimata K, Nakato H (2004) In vivo hyaluronan synthesis upon expression of the mammalian hyaluronan synthase gene in Drosophila. J Biol Chem 279 (18):18920–18925 14. Yuan H, Tank M, Alsofyani A, Shah N, Talati N, LoBello JC, Kim JR, Oonuki Y, de la Motte CA, Cowman MK (2013) Molecular mass dependence of hyaluronan detection by sandwich ELISA-like assay and membrane blotting using biotinylated hyaluronan binding protein. Glycobiology 23(11):1270–1280 15. Tolg C, Hamilton SR, Zalinska E, McCulloch L, Amin R, Akentieva N, Winnik F, Savani R, Bagli DJ, Luyt LG, Cowman MK, McCarthy JB, Turley EA (2012) A RHAMM mimetic peptide blocks hyaluronan signaling and reduces inflammation and fibrogenesis in excisional skin wounds. Am J Pathol 181(4):1250–1270 16. Yuan H, Amin R, Ye X, de la Motte CA, Cowman MK (2015) Determination of hyaluronan molecular mass distribution in human breast milk. Anal Biochem 474:78–88 17. Tolg C, Yuan H, Flynn SM, Basu K, Ma J, Tse KCK, Kowalska B, Vulkanesku D, Cowman MK, McCarthy JB, Turley EA (2017) Hyaluronan modulates growth factor induced mammary gland branching in a size dependent manner. Matrix Biol 63:117–132 18. Yingsung W, Zhuo L, Morgelin M, Yoneda M, Kida D, Watanabe H, Ishiguro N, Iwata H, Kimata K (2003) Molecular heterogeneity of the SHAP-hyaluronan complex. Isolation and characterization of the complex in synovial fluid from patients with rheumatoid arthritis. J Biol Chem 278(35):32710–32718 19. He H, Li W, Tseng DY, Zhang S, Chen SY, Day AJ, Tseng SC (2009) Biochemical characterization and function of complexes formed by hyaluronan and the heavy chains of interalpha-inhibitor (HC*HA) purified from extracts of human amniotic membrane. J Biol Chem 284(30):20136–20146

Chapter 9 Hyaluronan Pericellular Matrix: Particle Exclusion Assay Melanie A. Simpson Abstract Particle exclusion assays are used to visualize pericellular envelopes with a high content of hyaluronan. Pericellular hyaluronan is basally abundant in certain cell types while in others it is deposited in a highly dynamic manner in response to specific conditions and its presence may indicate cellular status. This assay, described here, is a quick semiquantitative approach to detecting pericellular hyaluronan using the hyaluronan-binding proteoglycan, aggrecan, to stabilize and amplify the surface matrix. Hyaluronan matrix can then be observed and quantified by microscopic image analysis of clear zones around individual cells, from which exogenously added fixed red blood cell particles are excluded. Key words Hyaluronan, Aggrecan, Pericellular matrix, Particle exclusion, Immunofluorescence

1

Introduction The particle exclusion assay has been a gold standard for pericellular matrix detection since first reported in 1968 by Clarris and Fraser [1]. In their report, the assay was developed for use in visualizing hyaluronidase-sensitive pericellular “zones of exclusion” that were individual properties of synovial cell strains and primary human embryonic fibroblasts. Subsequently, this assay has been used in many studies to detect pericellular matrices [2–6], confirm their hyaluronan (HA) content, characterize and compare the thickness of the HA envelopes [7–10], and examine the dynamics of HA matrix formation [11–16]. Such matrices are now known to be associated basally with specific cell types such as chondrocytes [17], with proliferating cells of many types [18, 19], on circulating cells of the innate immune system [20], and pathologically in response to pro-inflammatory signals [6, 19, 21]. HA matrices have also been associated with increased metastatic potential of tumor cells derived from multiple solid and blood-borne cancer types [22–27]. The particle exclusion assay capitalizes on the high molecular mass and hygroscopic properties of HA, which when encapsulating

Davide Vigetti and Achilleas D. Theocharis (eds.), The Extracellular Matrix: Methods and Protocols, Methods in Molecular Biology, vol. 1952, https://doi.org/10.1007/978-1-4939-9133-4_9, © Springer Science+Business Media, LLC, part of Springer Nature 2019

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a cell physically barricades the cellular boundary, thereby excluding small exogenously added particles. This is easily visualized by light microscopy. The areas of the matrix and cell boundaries can be measured and plotted as an index of average thickness. Further stabilization of an existing pericellular matrix can be achieved by the addition of an aggregating proteoglycan, aggrecan, which binds HA in the matrix and forms noncovalent cross-links that effectively augment the HA envelope. Some key limitations of the particle exclusion assay are the lack of molecular detail, semiquantitative nature of the assay, dependence on aggrecan addition to some cell types, low precision in comparing matrix sizes between cells, and two-dimensional context of the assay. Nonetheless, presence or absence of a HA matrix can be determined and further examined with fluorescent reporters, immunohistochemistry, flow cytometry, and other approaches. Here, we describe detailed protocols for detection and quantification of pericellular HA matrices and their thickness, as well as the validation of the assay using a molecular approach similar to immunofluorescence.

2 2.1

Materials Cell Culture

1. Minimal essential medium (MEM) or appropriate media containing 10% fetal bovine serum (FBS) (the growing medium depends on the cell type); filter sterilize if necessary, and store at 4  C. 2. Multiwell tissue culture plates, 48-well recommended.

2.2 Pericellular Matrix Visualization by Particle Exclusion

1. 1 MEM or appropriate media without phenol red (PRF MEM). 2. 1 MEM or appropriate media without phenol red, containing 0.1% BSA (PRF MEM/BSA): Add 0.1 g BSA to 100 mL PRF MEM. Use within 48 h or filter sterilize for longer term storage. Store at 4  C. 3. Sodium acetate buffer: 0.1 M Sodium acetate, pH 5.6, with 50 mM NaCl. Prepare a stock of 1 M sodium acetate, pH 5.6 in ultrapure H2O. Prepare a stock of 5 M NaCl in ultrapure H2O. Use 10 mL of each of these two stock solutions added to 80 mL of ultrapure H2O to prepare the final buffer. 4. Hyaluronidase from Streptomyces hyalurolyticus (Sigma H1136, also available from Fisher, EMD Millipore): Available lyophilized in vials of 300+ units. Dissolve 1 vial in 1 mL of sodium acetate buffer and aliquot for storage at 20  C. Use at a final concentration of 16 units/mL in 0.3 mL per well of a 48-well plate. 5. Aggrecan (Sigma A1960): Available lyophilized. Reconstitute 1 mg in 0.5 mL ultrapure H2O.

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6. Glutaraldehyde-fixed sheep red blood cells (Sigma R3378): Available lyophilized. Reconstitute in PBS, pH 7.2, as instructed by the vendor. Wash before using and resuspend at a final concentration of 5  108 per mL in PRF MEM. 2.3 Hyaluronan Matrix Detection by Fluorescence Imaging

1. PBS: Phosphate-buffered saline composed as 137 mM NaCl, 2.7 mM KCl, 4.3 mM Na2HPO4, 1.47 mM KH2PO4. Adjust to a final pH of 7.4. 2. PBS/4% paraformaldehyde (available from Fisher). 3. PBS/BSA: Add BSA to a final quantity of 0.1% (w/v) by using 0.1 mg of BSA per 100 mL PBS. 4. Hyaluronidase from Streptomyces h. (Sigma H1136 also Fisher, EMD Millipore), see Subheading 2.2, step 4, above. 5. Sodium acetate buffer: 0.1 M Sodium acetate, pH 5.6, with 50 mM NaCl, see Subheading 2.2, step 3, above. 6. Biotinylated hyaluronan-binding protein (bHABP): Available lyophilized from Millipore Sigma; dissolve 1 vial (50 μg) in 50 μL of ultrapure H2O for a 1 mg/mL stock. Use at 1:500 dilution in PBS containing 5% FBS. 7. Streptavidin-Alexa Fluor 568: Available from Thermo Fisher Scientific; use at 1:3000 dilution in PBS/BSA. 8. Glass coverslips, 12 mm diameter, 1.5 thickness. 9. Fluoromount-G (Thermo Fisher Scientific). 10. Clear nail polish (we routinely use Wet&Wild clear nail polish).

3

Methods

3.1 Particle Exclusion Assay

1. Seed cells in wells on a 48-well plate (depending on cell size and proliferation rate, seed 2000–5000 cells per well, see Note 1). Incubate overnight at 37  C and 5% CO2. 2. If using aggrecan to augment the HA matrix, prepare aggrecan solution (see Notes 2 and 3). Dilute aggrecan 1:10 from a 2 mg/mL stock in PRF MEM/BSA to make a final concentration of 0.2 mg/mL. Prepare enough diluted aggrecan to deliver 0.3 mL to each well being tested. 3. Wash all wells once with 0.5 mL of PRF MEM/BSA (see Note 4). 4. Treat experimental wells with 0.3 mL of Streptomyces hyaluronidase (16 units/mL, see Notes 5 and 6). For the working dilution, use 16 μL of a 300 unit/mL stock in 1:1 PRF MEM + sodium acetate buffer. Treat control wells with 0.3 mL per well of MEM with an equal ratio of PRF MEM and sodium acetate buffer (but without hyaluronidase).

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Fig. 1 Pericellular HA matrices revealed by particle exclusion assay. Human tumor cell lines were cultured in serum-containing media to subconfluence. Media were removed and replaced with sodium acetate assay buffer alone (a) or containing Streptomyces hyaluronidase (b). Cells were then washed and incubated for 90 min with aggrecan at 37  C, followed by addition of fixed sheep red blood cells. After allowing 15 min for the particles to settle, individual cells were imaged at 400 magnification

5. Incubate the plate at 37  C and 5% CO2 for 25 min and then wash wells once with 0.5 mL of PRF MEM/BSA. 6. If using aggrecan, add 0.3 mL of aggrecan solution to each well and incubate the plate at 37  C and 5% CO2 for 90 min. 7. Wash fixed red blood cells with 2 10 mL of PRF MEM and resuspend at 5  108/mL (see Note 7). 8. Wash all wells 2 with 0.5 mL of PRF MEM. 9. Add 0.3 mL of fixed red blood cell suspension to all wells. 10. Incubate plate at 37  C and 5% CO2 for 15–20 min to allow red blood cells to settle. 11. Examine cells with an inverted light microscope using phase contrast (see Note 8). Capture 15–20 individual images per condition at 400 magnification for Image J analysis (Fig. 1). 12. Analyze individual images in Image J by tracing matrix boundary and cell boundary to obtain areas of each. Plot as matrix:cell area ratio to compare matrix thickness (Fig. 2). 3.2 HA Matrix Validation by Fluorescence Detection

1. Culture cells overnight on coverslips. 2. Fix cells in PBS/4% paraformaldehyde for 10 min. 3. Wash 2 with 1 mL PBS. 4. For HA specificity validation, incubate one coverslip per condition in Streptomyces hyaluronidase and one coverslip in sodium acetate buffer.

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Fig. 2 Quantification of HA matrix thickness. Images of individual or small clusters of cells were captured at 400 magnification following the particle exclusion assay. Boundaries of cells (traced in black on the image at left) and boundaries of their corresponding exclusion zones (traced in white) were mapped for 20 cells per group using NIH Image J. From these tracings, respective matrix and cell areas were calculated and plotted as a ratio to compare pericellular matrix thickness. Hyaluronidase-treated cells are used as a control for HA content of the matrix and are expected to lack a matrix almost completely, which is indicated by a matrix-to-cell ratio that is close to 1:1

5. Prepare biotinylated HABP. Use 2 μg/mL with a volume of 25 μL per coverslip (12 mm diameter), diluted in PBS containing 5% FBS. 6. Incubate coverslip face down on droplet of bHABP for 60 min in a humidified Petri dish (see Note 9). 7. Wash coverslips gently with 2 1 mL of PBS. 8. Incubate with streptavidin-Alexa Fluor 568 (1:3000 in PBS / BSA) for 30 min at room temperature in a humidified Petri dish. 9. Wash 3 1–2 mL with PBS and mount with Fluoromount-G (see Note 10). 10. Seal with Wet&Wild clear nail polish (or other clear nail polish, see Note 10). 11. Image at 400 magnification by fluorescence confocal or epifluorescence microscopy (Fig. 3). 12. Perform quantitative fluorescence image analysis using NIH Image J.

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Fig. 3 HA pericellular visualization by fluorescence microscopy. Cells were cultured overnight on coverslips in serum-containing media. Prior to HA detection, cells were fixed with paraformaldehyde. Coverslips were washed and incubated in the absence (a) or presence (b) of Streptomyces hyaluronidase. After washing, cellsurface HA was probed by incubation with biotinylated HABP for 60 min followed by streptavidin-Alexa Fluor 568 for 30 min. Images were captured by fluorescence confocal microscopy

4

Notes 1. Cell density: The cells should be sparse with ample room in between individual cells. To obtain the ideal density for imaging and analysis of matrix thickness, it is helpful to pretest several seeding densities for each different cell type. 2. Phenol red-free media are recommended for better visualization and imaging of the pericellular matrices. However, phenol red-containing media can also be used. 3. Aggrecan may require time to dissolve homogeneously, which is critical for interpretable results. Warming the aggrecan solution in a 37  C water bath for 1–2 h is recommended. Inspect the solution visually for appearance of inhomogeneity. 4. Wash cells very gently at all steps. Cells are often poorly adherent and matrices may be disturbed by pipetting. 5. Streptomyces hyaluronidase is recommended for the confirmation of matrix HA content because this is the only hyaluronidase that is specific for HA as a substrate without acting on other GAGs that may be present. 6. The stock Streptomyces hyaluronidase is resuspended to 300 units/mL with sodium acetate buffer (stable for about a week at 4  C; aliquot and store at 20  C for longer term use) and then diluted to final concentration in MEM/acetate buffer. Although the slightly acidic pH of the acetate buffer is

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required for optimal activity of the enzyme, many cell types do not tolerate the pure acetate buffer. Adjust the MEM ratio as appropriate for the cells. 7. It is very important to wash the fixed red blood cells before using. This will help prevent them from auto-aggregating and will disperse residue from lysed cells. Following the washes, resuspend the red blood cells in the volume needed to deliver 0.3 mL of red blood cell suspension to each well of the assay plate. 8. Do not allow plates to sit longer than about 30–40 min. Results will be compromised by increased settling of RBC onto the cells of interest. This may or may not obscure the halos, but can also disrupt the visual clarity because of the adherence of RBC to the cell bodies. 9. Humidified Petri dish: Place a moist filter or paper towel flat in the bottom of a 10 cm petri dish to preserve moisture in the bHABP droplet during incubations. The 25 μL droplet is pipetted on a glass slide and the coverslip placed face down over it, allowing a minimal volume of bHABP to be used. The slide is placed flat (coverslip up) on the paper towel and the Petri dish covered during incubation. 10. Use of these additive and sealant reagents promotes longevity of the fluorescence signal. References 1. Clarris BJ, Fraser JR (1968) On the pericellular zone of some mammalian cells in vitro. Exp Cell Res 49:181–193 2. Heldin P, Pertoft H (1993) Synthesis and assembly of the hyaluronan-containing coats around normal human mesothelial cells. Exp Cell Res 208:422–429 3. Knudson W, Knudson CB (1991) Assembly of a chondrocyte-like pericellular matrix on non-chondrogenic cells. Role of the cell surface hyaluronan receptors in the assembly of a pericellular matrix. J Cell Sci 99:227–235 4. McCarthy MT, Toole BP (1989) Membraneassociated hyaluronate-binding activity of chondrosarcoma chondrocytes. J Cell Physiol 141:191–202 5. Moriarty KP, Crombleholme TM, Kerry Gallivan E, O’Donnell C (1996) Hyaluronic acid-dependent pericellular matrices in fetal fibroblasts: implication for scar-free wound repair. Wound Repair Regen 4(3):346–352. https://doi.org/10.1046/j.1524-475X. 1996.40311.x

6. Toole BP (1997) Hyaluronan in morphogenesis. J Int Med 242:35–40 7. Bullard KM, Kim HR, Wheeler MA, Wilson CM, Neudauer CL, Simpson MA, McCarthy JB (2003) Hyaluronan synthase-3 is upregulated in metastatic colon carcinoma cells and manipulation of expression alters matrix retention and cellular growth. Int J Cancer 107:739–746 8. Knudson CB, Nofal GA, Pamintuan L, Aguiar DJ (1999) The chondrocyte pericellular matrix: a model for hyaluronan-mediated cellmatrix interactions. Biochem Soc Trans 27:142–147 9. Laurent TC, Fraser JR (1992) Hyaluronan. FASEB J 6:2397–2404 10. Simpson MA, Reiland J, Burger SR, Furcht LT, Spicer AP, Oegema TR Jr, McCarthy JB (2001) Hyaluronan synthase elevation in metastatic prostate carcinoma cells correlates with hyaluronan surface retention, a prerequisite for rapid adhesion to bone marrow endothelial cells. J Biol Chem 276:17949–17957

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11. Evanko SP, Angello JC, Wight TN (1999) Formation of hyaluronan- and versican-rich pericellular matrix is required for proliferation and migration of vascular smooth muscle cells. Arterioscler Thromb Vasc Biol 19:1004–1013 12. King A, Balaji S, Le LD, Marsh E, Crombleholme TM, Keswani SG (2013) Interleukin-10 regulates fetal extracellular matrix hyaluronan production. J Pediatr Surg 48:1211–1217 13. Maleski M, Hockfield S (1997) Glial cells assemble hyaluronan-based pericellular matrices in vitro. Glia 20:193–202 14. Nishida Y, Knudson CB, Knudson W (2003) Extracellular matrix recovery by human articular chondrocytes after treatment with hyaluronan hexasaccharides or Streptomyces hyaluronidase. Mod Rheumatol 13:62–68 15. Nishida Y, Knudson CB, Kuettner KE, Knudson W (2000) Osteogenic protein-1 promotes the synthesis and retention of extracellular matrix within bovine articular cartilage and chondrocyte cultures. Osteoarthr Cartil 8:127–136 16. Simpson MA, Wilson CM, Furcht LT, Spicer AP, Oegema TR Jr, McCarthy JB (2002) Manipulation of hyaluronan synthase expression in prostate adenocarcinoma cells alters pericellular matrix retention and adhesion to bone marrow endothelial cells. J Biol Chem 277:10050–10057 17. Knudson W, Ishizuka S, Terabe K, Askew EB, Knudson CB (2018) The pericellular hyaluronan of articular chondrocytes. Matrix biol. https://doi.org/10.1016/j.matbio.2018.02. 005 18. Evanko SP, Tammi MI, Tammi RH, Wight TN (2007) Hyaluronan-dependent pericellular matrix. Adv Drug Deliv Rev 59:1351–1365 19. Toole BP (2004) Hyaluronan: from extracellular glue to pericellular cue. Nat Rev Cancer 4:528–539

20. Lee-Sayer SS, Dong Y, Arif AA, Olsson M, Brown KL, Johnson P (2015) The where, when, how, and why of hyaluronan binding by immune cells. Front Immunol 6:150 21. Nishida Y, D’Souza AL, Thonar EJ, Knudson W (2000) Stimulation of hyaluronan metabolism by interleukin-1alpha in human articular cartilage. Arthritis Rheum 43:1315–1326 22. Kim HR, Wheeler MA, Wilson CM, Iida J, Eng D, Simpson MA, McCarthy JB, Bullard KM (2004) Hyaluronan facilitates invasion of colon carcinoma cells in vitro via interaction with CD44. Cancer Res 64:4569–4576 23. Lessan K, Aguiar DJ, Oegema T, Siebenson L, Skubitz AP (1999) CD44 and beta1 integrin mediate ovarian carcinoma cell adhesion to peritoneal mesothelial cells. Am J Pathol 154:1525–1537 24. Nakazawa H, Yoshihara S, Kudo D, Morohashi H, Kakizaki I, Kon A, Takagaki K, Sasaki M (2006) 4-methylumbelliferone, a hyaluronan synthase suppressor, enhances the anticancer activity of gemcitabine in human pancreatic cancer cells. Cancer Chemother Pharmacol 57:165–170 25. Ricciardelli C, Russell DL, Ween MP, Mayne K, Suwiwat S, Byers S, Marshall VR, Tilley WD, Horsfall DJ (2007) Formation of hyaluronanand versican-rich pericellular matrix by prostate cancer cells promotes cell motility. J Biol Chem 282:10814–10825 26. Simpson MA (2006) Concurrent expression of hyaluronan biosynthetic and processing enzymes promotes growth and vascularization of prostate tumors in mice. Am J Pathol 169:247–257 27. Tammi MI, Oikari S, Pasonen-Seppanen S, Rilla K, Auvinen P, Tammi RH (2018) Activated hyaluronan metabolism in the tumor matrix causes and consequences. Matrix Biol. https:// doi.org/10.1016/j.matbio.2018.04.012

Chapter 10 Determination of Cell-Surface Hyaluronan Through Flow Cytometry Daiana L. Vitale, Fiorella M. Spinelli, and Laura Alaniz Abstract Hyaluronan is the major glycosaminoglycan present in the extracellular matrix of several cell types; its synthesis occurs on the cellular plasma membrane. Variations in its expression are related to alterations in cell proliferation, adhesion, and migration. It is able to interact with different binding proteins called hyaladherins, which can be conjugated to different fluorochromes and analyzed by flow cytometry. Key words Extracellular matrix, Hyaluronan, Hyaluronan-binding proteins, Flow cytometry

1

Introduction The extracellular matrix (ECM) is the noncellular component present within all tissues and organs. The ECM provides not only essential physical support for cellular constituents but also initiates crucial biochemical processes that are required for tissue morphogenesis, differentiation, and homeostasis. Fundamentally, the ECM is composed of proteins and polysaccharides, and each tissue has a unique ECM composition and topology that is generated during tissue development [1]. Among most abundant components of ECM are glycosaminoglycans (GAGs): long unbranched polysaccharides, composed of repeated units of disaccharides of uronic acid and amino sugars. Hyaluronan (HA) is the major GAG found in all types of mammalian ECM. HA biosynthesis occurs on the cellular plasma membrane by means of three HA synthase isoenzymes (HAS-1, HAS-2, HAS-3) [2]. These enzymes are integral membrane proteins that catalyze the alternate addition of glucuronic acid and N-acetyl glucosamine from their UDP derivatives to a growing HA polymer, continuously translocated through the plasma membrane into ECM [3]. Besides having structural properties, HA is able to modulate several biological processes through the interaction with different proteins called hyaladherins or

Davide Vigetti and Achilleas D. Theocharis (eds.), The Extracellular Matrix: Methods and Protocols, Methods in Molecular Biology, vol. 1952, https://doi.org/10.1007/978-1-4939-9133-4_10, © Springer Science+Business Media, LLC, part of Springer Nature 2019

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hyaluronan-binding proteins (HABP), which include the main membrane receptor CD44, within others such as RHAMM, TSG-6, and versican [4]. The interaction between HA and CD44 mediates different physiological processes such as adhesion, proliferation, and cell migration. However, it has shown that abnormal production of HA promotes also pathological processes such as inflammation [5] or tumor progression [6]. Flow cytometry is a well-established technique mainly used in clinical diagnostics and biomedical research. It is designed to deliver cells in single file at the point of measurement. Light signals emitted from cells are collected and correlated to morphology, surface, and intracellular biomolecule expression [7]. Single-cell suspensions are required for flow cytometry assays. Thus, cells that grow in suspension are well suited for analysis by flow cytometry. Adherent cell lines and solid tissue samples, such as tumors, require processing into single-cell suspensions before being analyzed. Since HA synthesis occurs on the cell plasma membrane, it is possible to use flow cytometry to determine the relative amount of HA present on the cell surface, using different HABPs conjugated to biotin or fluorescence dye. In fact, we are using biotinconjugated HABPs that bind to fluorescent-streptavidin dye in order to amplify the intensity of the signal.

2 2.1

Materials Cell Harvesting

1. Phosphate-buffered saline (PBS; 10 1 L): 1.37 M NaCl, 0.027 M KCl, 0.255 M Na2HPO4, 0.018 M KH2PO4, pH 7.4. Once the 10 stock solution is prepared, dilute 100 mL in 900 mL of distilled water to obtain ready-to-use 1 solution. Store at 4  C. 2. Trypsin-EDTA (1) solution: 0.25% Trypsin and 0.1% EDTA in PBS or rubber-tipped cell scraper. 3. 0.4% Trypan Blue solution in PBS.

2.2 Sample Preparation for Labeling

1. FACS buffer: Fetal bovine serum (FBS) 1% and 0.05% sodium azide in PBS. 2. Biotinylated-HABP (Cat # 385911, Calbiochem). 3. Fluorescent-labeled streptavidin. 4. Hyaluronidase (300 U/mL, Cat # H3506, Sigma Aldrich). 5. Paraformaldehyde (PFA): 4% Solution in ultrapure water (see Note 1).

Determination of Cell-Surface Hyaluronan

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Methods

3.1 Adherent Cell Line Processing

1. Discard supernatant and wash cells with warm PBS. 2. Detach cells from the plate using a cell scraper and collect cells with 1 mL of cold PBS or treating with trypsin (see Note 2). 3. Wash cells by centrifugation for 5 min at 380  g and resuspend in cold PBS (see Note 3). 4. Check cell viability by Trypan Blue dye exclusion (see Note 4). 5. Count cells using a hemocytometer. 6. Resuspend in PBS at 5  105 to 1  106 cells per 100 μL and keep on ice.

3.2 Cell Line Suspension Processing

1. Decant cells by centrifugation for 5 min at 380  g at room temperature (RT). 2. Check cell viability by Trypan Blue dye exclusion (see Note 4). 3. Count cells using a hemocytometer. 4. Wash cells centrifuging for 5 min at 380  g with PBS at 4  C and resuspend in cold PBS (see Note 3). 5. Resuspend in PBS at 5  105 to 1  106 cells per 100 μL and keep on ice.

3.3

Cell Staining

Unless otherwise noted, the procedure is carried out on ice (Fig. 1 near here). 1. Consider four different controls and in addition the samples to perform the following essay (see Note 5): (a) Control 1 tube: autofluorescence control (unstained cells) (see Note 6). (b) Control 2 tube: control of stained cells only with biotinylated-HABP. (c) Control 3 tube: control of stained cells only with fluorescent-labeled streptavidin. (d) Control 4 tube: hyaluronidase treatment (negative control) (see Note 7). 2. Add 5 μg of biotinylated-HABP for each sample, and control tube 2 and tube 4 (this latter previously treated with hyaluronidase). 3. Incubate for 60 min at 2–8  C or on ice. It is not necessary to protect from light. 4. Wash cells by adding 1 mL of FACS buffer and centrifuging for 5 min at min at 380  g at 4  C. 5. Discard supernatant and resuspend in 100 μL of FACS buffer.

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Fig. 1 General procedure for detection of cell-surface hyaluronan through flow cytometry

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6. Add fluorescent-labeled streptavidin (include control tube 3) at 1:100 dilution (see Note 8). 7. Incubate for at least 45 min at 2–8  C or on ice. Protect from light. 8. Take all controls (1–4, and) and samples and wash twice cells by adding 1 mL of FACS buffer and centrifuging for 5 min at min at 380  g at 4  C. 9. Discard supernatant and resuspend all samples in an appropriate volume (250–400 μL) of FACS buffer (see Note 9). 10. For a later analysis (within a week), when a cytometer is available, we recommend to fix the cells as described in Subheading 3.4 (see Note 10). 3.4

Fixation

1. Suspend the cells in 1% PFA-PBS (see Note 11). 2. Store cells at 2–8  C, approximately for 1 week. Protect from light.

4

Notes 1. Store at 4  C and protect from light. 2. Trypsin treatment should not exceed 3–5 min at 37  C. Longer treatments may be prejudicial to analyze certain components of the cytoplasmic membrane. Make sure to inactivate trypsin adding the double amount of trypsin volume of culture medium supplemented with 10% FBS before centrifuge. 3. Protein-binding kinetics are temperature dependent. We recommend staining with ice-cold reagents/solutions, and at 4  C, as low temperature prevents the modulation and internalization of surface antigens. Staining on ice may require longer incubation times. 4. Cell viability must be around of 90%, to avoid subsequent problems during flow cytometry assay. 5. Internal controls are important to ensure the authenticity of obtained data and to avoid nonspecific fluorescent signal. Make sure that the fluorescence detection parameters are correctly adjusted during flow cytometer acquisition. 6. It is recommended to use more cells for autofluorescence control (tube 1) with respect to other samples since it will be used for the correct setting of FACS instrument during analyses. Control 1 tube (autofluorescence control) with unstained cells establishes the levels of background fluorescence during flow cytometer acquisition.

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7. For control number 4, treat cells with 2–3 U/mL of hyaluronidase for 1 h at 37  C. 8. At the first time, it is necessary to check different concentrations of this reagent (fluorescent-labeled streptavidin) in order to find the most suitable concentration according to your products (for example 1, 5, 10 μg; 1:100, 1:200, 1:500 dilutions, see the manufacturer’s recommendations). 9. In general, cells can be resuspended in 250–400 μL of FACS buffer. It is recommended to maintain a concentrated suspension of cells to facilitate the analysis. 10. We recommend analysis on the same day. The cells should be fixed for extending storage as well as for greater flexibility in planning time on the cytometer. 11. This procedure preserves cells for at least 1–2 weeks. It is recommended to use single fluorophores, such as Alexa Fluor488 or FITC. Since the brightness of tandem dyes might be reduced by the fixation process as short as possible. Fixation will stabilize the light scatter and inactivate most biohazardous agents. Controls will require fixation using the same procedure. IMPORTANT: Cells should not be fixed if they need to remain viable. References 1. Jarvelainen H, Sainio A, Koulu M, Wight TN, Penttinen R (2009) Extracellular matrix molecules: potential targets in pharmacotherapy. Pharmacol Rev 61(2):198–223 2. Vigetti D, Karousou E, Viola M, Deleonibus S, De Luca G, Passi A (2014) Hyaluronan: biosynthesis and signaling. Biochim Biophys Acta 1840 (8):2452–2459 3. Weigel PH, Hascall VC, Tammi M (1997) Hyaluronan synthases. J Biol Chem 272 (22):13997–14000 4. Day AJ, Prestwich GD (2002) Hyaluronanbinding proteins: tying up the giant. J Biol Chem 277(7):4585–4588

5. Grivennikov SI, Greten FR, Karin M (2010) Immunity, inflammation, and cancer. Cell 140 (6):883–899 6. Alaniz L, Rizzo M, Malvicini M, Jaunarena J, Avella D, Atorrasagasti C, Aquino JB, Garcia M, Matar P, Silva M, Mazzolini G (2009) Low molecular weight hyaluronan inhibits colorectal carcinoma growth by decreasing tumor cell proliferation and stimulating immune response. Cancer Lett 278(1):9–16 7. Ibrahim SF, van den Engh G (2007) Flow cytometry and cell sorting. Adv Biochem Eng Biotechnol 106:19–39

Chapter 11 Method for Detecting Hyaluronan in Isolated Myenteric Plexus Ganglia of Adult Rat Small Intestine Michela Bistoletti, Paola Moretto, and Cristina Giaroni Abstract The cellular components of the enteric nervous system (ENS), namely enteric neurons and glia, display plasticity and respond to environmental cues deriving from growth factors, extracellular matrix (ECM) molecules, and cell-surface molecules, both in physiological and pathological conditions. ECM, in particular, provides an important framework for the enteric microenvironment and influences the homeostasis of myenteric neuronal circuitries. Isolation of pure myenteric plexus preparations from adult tissue permits to investigate changes in the ENS involving specific ECM, such as hyaluronan. This approach is based upon the possibility to isolate myenteric ganglia from the intestinal wall of either adult animals or humans, after microdissection and subsequent enzymatic digestion of the tissue. Enteric ganglia are free of connective tissue, extracellular collagen, and blood vessels, and thus treatment of intact intestinal segments with highly purified collagenases permits ganglia isolation from the surrounding smooth muscle cells. In this chapter, we describe methods for visualizing HA in isolated primary cultures of adult rat small intestine myenteric ganglia. Key words Enteric nervous system (ENS), Myenteric ganglia, Hyaluronan, Extracellular matrix, Hyaluronan-binding protein (HABP)

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Introduction The enteric nervous system (ENS) is a complex and highly integrated network of ganglia and interconnecting fibers forming two main ganglionated plexuses, the submucosal and myenteric plexus, responsible for regulating the digestive functions [1, 2]. The cellular components of the ENS, namely enteric neurons and glia, display plasticity and respond to environmental cues deriving from growth factors, extracellular matrix molecules (ECM), and cell-surface molecules both in physiological conditions (i.e., during development) and in pathological conditions (i.e., during inflammation, ischemia/reperfusion damage) [3–7]. ECM, in particular, provides an important framework for the enteric microenvironment and influences the homeostasis of

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myenteric neuronal circuitries. Recent evidence has been provided to show that myenteric neurons produce hyaluronan (HA), which retains a homeostatic role by contributing to the formation of an extracellular matrix basal membrane enveloping the surface of myenteric ganglia as well as a perineuronal net surrounding myenteric neurons [8]. This well-organized HA structure is highly altered after an experimentally induced colitis, suggesting that the glycosaminoglycan may participate in myenteric neuron derangement underlying motility changes during intestinal inflammation. Studies aiming at elucidating the factors that influence the ENS functions, such as neurotransmitter pathways, signaling cascades, and ECM, are mainly carried out with classic techniques resorting to intact segments of intestinal preparations with embedded myenteric plexus [9–11]. However, these techniques do not allow to investigate molecular pathways specifically influencing enteric neuron or glia, since they consist of a mixture of cells. Isolation of pure myenteric plexus preparations from adult tissue may overcome this limitation allowing to study molecular changes involving the ENS both in physiological and in pathological conditions [12]. This approach is based on the possibility to isolate myenteric ganglia from the intestinal wall of either adult animals or humans, after microdissection and subsequent enzymatic digestion of the tissue [13, 14]. Enteric ganglia are free of connective tissue, extracellular collagen, and blood vessels [15, 16], and thus treatment of intact intestinal segments with highly purified collagenases permits ganglia isolation from the surrounding smooth muscle cells [13, 14]. The present technique can be used to perform a variety of types of research including electrophysiological approaches, live/dead assays, calcium imaging, ROS production, release of inflammatory modulators, electron microscopy, and fluorescence microscopy. In the following paragraphs we describe a method for visualizing HA in isolated primary cultures of myenteric ganglia resorting to a fluorescently labeled HA-binding protein (HABP) as a marker to detect the glycosaminoglycan by confocal microscopy. HABP recognizes HA saccharidic sequences and is able to localize HA in tissues by streptavidin conjugation with an appropriate fluorophore [17].

2

Materials 1. Round coverslips (12 mm in diameter, 0.13–0.17 mm in thickness, autoclaved). 2. Plastic 24-well plate. 3. Scissors. 4. Microdissection scissors.

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5. Forceps. 6. Stereomicroscope. 7. Laminar flow hood. 8. Petri dish covered with Sylgard (Dow Corning, Seneffe, Belgium). 9. Entomology pins. 10. 15 mL Conical tubes. 11. Petri dish (35 mm in diameter). 12. Rotator or shaker. 13. Centrifuge. 14. Incubator (37
C, 5% CO2). 15. Pipettes and sterile tips. 16. Becher. 15. Dark humid chamber. 2.1

Reagents

1. Phosphate-buffered saline (PBS), composition: 0.14 M NaCl, 0.003 M KCl, 0.015 M Na2HPO4, 0.0015 M KH2PO4, pH 7.4. 2. Ca2+- and Mg2+-free Hanks’ Balanced Salt Solution (HBSS, Thermo Fisher, Milan, Italy). 3. 100 ng/L Poly-L-lysine (Sigma Aldrich, Milan, Italy) in HBSS. 4. Dulbecco’s modified Eagle’s medium (DMEM, Thermo Scientific, Milan, Italy). 5. Fetal bovine serum (FBS, Euroclone, Milan Italy). 6. Myenteric ganglia complete culture medium: DMEM supplemented with 10% FBS, 0.5% glutamine, 1% penicillin/streptomycin, 50 μg/mL gentamycin sulfate, 10 ng/mL glial cell linederived neurotrophic factor (GDNF, Sigma Aldrich). 7. Enzyme solution: HBSS containing 1.5 mg/mL collagenase Type I (from Clostridium histolyticum, Sigma Aldrich) and 1.25 mg/mL protease Type I (from bovine pancreas, Sigma Aldrich), 1 mg/mL DNase (from bovine pancreas, Sigma Aldrich). 8. 1.25 mg/mL Trypsin with 0.01% ethylenediaminetetraacetic acid (EDTA, Sigma Aldrich) pH 7.4. 9. Physiological Tyrode’s solution (composition: 137 mM NaCl; 2.68 mM KCl; 1.8 mM CaCl2·2H2O; 2 mM MgCl2; 0.47 mM NaH2PO4; 11.9 mM NaHCO3; 5.6 mM glucose). 10. PBS containing 4% formaldehyde: Prepare immediately before use.

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11. Blocking buffer: PBS containing 0.1% Triton X-100 (Sigma Aldrich) and 5% normal horse serum (NHS, Euroclone). 12. Primary antibodies: Biotin-labeled HA-binding protein (HABP) (Seikagaku, AMS Biotechnology Europe, UK); biotin-conjugated HuC/D mouse anti-human primary antibody (Thermo Fisher); S-100β rabbit anti-cow primary antibody (Dako, Denmark). 13. Secondary antibodies: Cy3-conjugated streptavidin (Molecular Probes, Invitrogen, Carlsbad, CA), FITC-conjugated streptavidin (Molecular Probes, Invitrogen, Carlsbad, CA), F(ab0 )2 anti-rabbit IgG (H+L) biotin secondary antibody (Caltag laboratories, Inc., Baltimore, USA). 14. Mounting medium with DAPI (Vectashield®, Vector Lab., Burlingame, CA).

3

Methods

3.1 Preparation of Intestinal Myenteric Ganglia Cultures and Fixation

Day 1:

1. Prepare 1 day before myenteric ganglia isolation (see Note 1). 2. Put 12 mm autoclaved coverslips in a 24-well plate; wash the wells with PBS two times. 3. Remove PBS and add 200 μL poly-L-lysine solution per coverslip. Incubate at room temperature for at least 3 h. 4. Remove the poly-L-lysine solution and wash the coverslips with PBS three times. 5. Add 1 mL of myenteric ganglia culture medium to each well and incubate the plate overnight at room temperature. Day 2:

6. Replace the medium with 1 mL of fresh culture medium per well. 7. Sacrifice one animal according to the protocols approved by the Animal Care and Use Ethics Committee of your institution. 8. Open the abdomen with scissors without injuring the bowel. 9. Remove immediately a 20 cm long segment of the small intestine, 5 cm oral to the ileocecal junction, and place it in a becher containing a physiological ice-cold Tyrode’s solution. Cut the intestinal segment into 2 cm long specimens and perfuse them with Tyrode’s solution, by means of a 2.5 mL syringe, in order to flush out feces. 10. Put the segment on a Sylgard-covered petri dish and wet it with HBBS + Hepes. Gently remove the mesentery with the excess

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of fat tissue from the outside of the intestine, under a stereomicroscope, by means of sterile microdissecting scissors. 11. Fix the intestinal segment, with the serosal side up, by means of entomology pins on Sylgard. Gently strip the longitudinal muscle with attached myenteric ganglia (LMMP) from the whole intestinal wall using fine sterile forceps (see Note 2). 12. Mechanically mince the LMMP segments with sterile microdissecting scissors. Collect the minced tissue in a 15 mL conical tube filled with 10 mL of pre-warmed enzyme solution. Incubate at 37
C for 25 min on a shaker or rotator at low speed. 13. Add 1 mL of the trypsin solution with EDTA, and incubate at 37
C for 20 min. 14. Add FBS (6% v/v). 15. Centrifuge the cell suspension at 1500  g for 7 min at 4
C. Remove the supernatant. Resuspend the cell pellet in 10 mL HBSS. 16. Repeat step 15 two times. 17. Resuspend the cell pellet in 4 mL of DMEM complete culture medium and transfer the suspension in a 35 mm petri dish. 18. Remove aggregates of smooth muscle cells from ganglia and collect them using a 200 μL pipette under a stereomicroscope (see Note 3). 19. Transfer isolated ganglia in 1 mL of DMEM complete culture medium and seed them on the poly-L-lysine-pre-coated glass coverslips into the 24-well plate (Fig. 1). 20. Incubate cells at 37
C in an incubator (37
C, 5% CO2) for 6 days, replacing the medium every 2 days. 21. On the 7th day, proceed for cell fixation by removing the medium from the wells and adding 500 μL of PBS containing 4% formaldehyde in each of them. Fix cells for 10 min at 37
C. 22. Remove the fixative solution. 23. Proceed with the immunofluorescence staining protocol as described in Subheading 3.2. The different phases of myenteric ganglia isolation are described in Fig. 1. 3.2 Immunofluorescence Staining

To visualize and localize hyaluronan in isolated primary cultures of myenteric ganglia we perform double labeling with HABP and primary antibodies either for the pan-neuronal marker HUC/D or for the enteric glial cell marker S100β, during consecutive incubation times.

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Fig. 1 Schematic representation of the myenteric ganglia dissection procedure. LMMP longitudinal muscle myenteric plexus

1. Wash preparations consisting of coverslips with attached myenteric ganglia for 5 min with 1 mL of PBS (1) on a rotator. Repeat two times. 2. Remove PBS (1) and add in each well 200 μL of blocking buffer. Incubate the plate for 1 h at room temperature to permeabilize the preparations and to block nonspecific binding sites (see Note 4). Gently remove the buffer. 3. Apply biotin-labeled HABP (5 μg/mL, see Note 5), and incubate overnight at 4
C. 4. Remove HABP and wash preparations three times with PBS (1), each for 5 min. 5. Apply a FITC conjugated-streptavidin complex (dilution 1:300). Incubate at RT for 2 h.

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Fig. 2 Phase micrographs of adult rat ileum cultured myenteric ganglia, at day 1 (a) and after 6 days of culture (b). Arrows indicate neurons. Bar: 25 μm

6. Remove the streptavidin complex and wash preparations three times with PBS (1), each for 5 min. 7. Apply either biotin-conjugated HuC/D or S100β primary antibody with a dilution of 1:100 and 1:200, respectively (see Note 5). Incubate overnight at 4
C. 8. Remove the antibody and wash with PBS (1) three times. 9. Apply either Cy3-conjugated-streptavidin complex (dilution 1:500) to reveal HUC/D labeling or a F(ab0 )2 anti-rabbit IgG biotin secondary antibody (1:300) followed by Cy3-conjugated streptavidin complex (dilution 1:500) to reveal S100β labeling (see Note 5). Incubate preparations for 1.5 h at RT. 10. Wash the preparations with PBS (1) three times, each of 5 min. 11. Remove the coverslips with cells from the wells using fine forceps, and mount them onto glass slides using a mounting medium with DAPI. 12. Analyze preparations by confocal microscopy (Figs. 2 and 3).

4

Notes 1. All steps have to be carried out under a laminar flow hood to maintain the sterility. 2. All incubations have to be carried out in a dark humid chamber to preserve the stability of antibodies. 3. This step has to be performed under a stereomicroscope to avoid the unintentional removal of already isolated plexus.

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Fig. 3 HA staining in primary cultures of rat small intestine isolated myenteric ganglia. (a–c) Confocal image showing co-localization of HABP with the pan neuronal marker HuC/D. Arrows indicate neurons. Bar: 25 μm. (d–f) Confocal image showing the absence of co-localization between HABP and the glial marker S100β. Arrowheads indicate a myenteric ganglion. Bar: 25 μm

4. Pay attention not to penetrate the gut mucosa during this step, thereby avoiding contaminations. 5. All dilutions for primary and secondary antibodies are made in blocking buffer. References 1. Furness JB, Callaghan BP, Rivera LR, Cho HJ (2014) The enteric nervous system and gastrointestinal innervation: integrated local and central control. Adv Exp Med Biol 817:39–71 2. Filpa V, Moro E, Protasoni M, Crema F, Frigo G et al (2016) Role of glutamatergic neurotransmission in the enteric nervous system and brain-gut axis in health and disease. Neuropharmacology 111:14–33 3. Giaroni C (2015) Purinergic signalling and development of the autonomic nervous system. Auton Neurosci 191:67–77 4. Lomax AE, Ferna´ndez E, Sharkey KA (2005) Plasticity of the enteric nervous system during intestinal inflammation. Neurogastroenterol Motil 17(1):4–15 5. Giaroni C, Marchet S, Carpanese E, Prandoni V, Oldrini R, Bartolini B et al (2013) Role of neuronal and inducible nitric oxide synthases in the guinea pig ileum

myenteric plexus during in vitro ischemia and reperfusion. Neurogastroenterol Motil 23: e114–e126 6. Giaroni C, Zanetti E, Giuliani D, Oldrini R, Marchet S, Moro E et al (2011) Protein kinase C modulates NMDA receptors in the myenteric plexus of the guinea pig ileum during in vitro ischemia and reperfusion. Neurogastroenterol Motil 23:e91–e103 7. Filpa V, Carpanese E, Marchet S, Pirrone C, Conti A et al (2017) Nitric oxide regulates homeoprotein OTX1 and OTX2 expression in the rat myenteric plexus after intestinal ischemia-reperfusion injury. Am J Physiol Gastrointest Liver Physiol 312:G374–G389 8. Filpa V, Bistoletti M, Caon I, Moro E, Grimaldi A et al (2017) Changes in hyaluronan deposition in the rat myenteric plexus after experimentally-induced colitis. Sci Rep 7:17644

Hyaluronan Detection in Isolated Myenteric Ganglia 9. Caputi V, Marsilio I, Filpa V, Cerantola S, Orso G et al (2017) Antibiotic-induced dysbiosis of the microbiota impairs gut neuromuscular function in juvenile mice. Br J Pharmacol 174 (20):3623–3639 10. Giaroni C, Zanetti E, Pascale A, Oldrini R, Canciani L et al (2009) Involvement of Ca2+dependent PKCs in the adaptive changes of mu-opioid pathways to sympathetic denervation in the guinea pig colon. Neurogastroenterol Motil 78(9):1233–1241 11. Filpa V, Carpanese E, Marchet S, Prandoni V, Moro E et al (2015) Interaction between NMDA glutamatergic and nitrergic enteric pathways during in vitro ischemia and reperfusion. Eur J Pharmacol 750:123–131 12. Carpanese E, Moretto P, Filpa V, Marchet S, Moro E et al (2014) Antagonism of ionotropic glutamate receptors attenuates chemical

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ischemia-induced injury in rat primary cultured myenteric ganglia. PLoS One 9(11):e113613 13. Jaeger CB (1995) Isolation of enteric ganglia from the myenteric plexus of adult rats. J Neural Transplant Plast 5:223–232 14. Sch€afer KH, Saffrey MJ, Burnstock G, MestresVentura P (1997) A new method for the isolation of myenteric plexus from the newborn rat gastrointestinal tract. Brain Res Brain Res Protoc 1:109–113 15. Gabella G (1979) Innervation of the gastrointestinal tract. Int Rev Cytol 59:129–193 16. Gershon MD (1981) The enteric nervous system. Annu Rev Neurosci 4:227–272 17. Raio L, Cromi A, Ghezzi F, Passi A, Karousou E et al (2005) Hyaluronan content of Wharton’s jelly in healthy and Down syndrome fetuses. Matrix Biol 24:166–174

Chapter 12 Analyzing Hyaluronidases in Biological Fluids Christos Velesiotis, Stella Vasileiou, and Demitrios H. Vynios Abstract Hyaluronidases are a group of enzymes responsible for the degradation of hyaluronan. They seem to be associated with a plethora of pathological conditions, as it has been showcased by numerous studies over the past years. The emerging role of hyaluronidases in various pathological states, especially cancer, is of a great interest. Thus, they are considered as important research targets. In this chapter the popular assay for hyaluronidase analysis in biological fluids is presented and discussed in detail. The assay is divided into two steps; the first is zymography that aims mainly to detect different hyaluronidase enzymes in a biological sample, and the second is the direct quantitative measurement of enzymatic activity by a microtiter plate assay. Both steps are characterized by high sensitivity, simplicity, and limited time consumption. Key words Hyaluronidase, Hyaluronan, Zymography, Substrate-electrophoresis, ELISA-like, Biotinavidin

1

Introduction Hyaluronidases (Hyals) are a class of degradative enzymes present in several toxins and venoms, helping their spreading in the body. Testicular Hyal, present in mammals in the sperm acrosome, is necessary for the fertilization of the ovum. Bacterial, invertebrate, and testicular Hyals have been extensively studied, so far. During the last 30 years the studies have focused on Hyals present in higher organisms, especially humans, since a connection was observed between Hyals and various pathological states, more characteristically in cancer [1–3]. Moreover, advances in the field of Hyals include evidence for the presence of Hyal and Hyal-like enzymes in various organisms and fluids [4–8] and production of recombinant Hyals, under the form of either the whole enzyme [9–11] or the catalytic domain [12]. Hyals predominantly degrade hyaluronan (HA), but they can also cleave chondroitin sulfate and chondroitin at a slower rate. Hyals are endoglycosidases, as they degrade the β-N-acetyl-D-glucosaminidic linkages in the HA polymer. In the human genome, six

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Hyal genes are present, namely HYAL-1, -2, and -3, clustered in the chromosome 3p21.3 locus, and HYAL-4, HYAL-P1, and PH-20, clustered in the chromosome 7q31.3 locus. It should be noted that HYAL-P1 is transcribed but not translated and HYAL-4 is rather a chondroitin-cleaving enzyme. According to their pH activity profiles, Hyals are divided into two categories: the acidic Hyals HYAL1, -2, and -3, which are active at acidic pH, and the neutral active Hyal PH-20 which has an activity profile in a broader pH range 3.0–9.0. The well-characterized mammalian Hyals are Hyal-1 and -2 and PH-20. Hyal-1 is a lysosomal enzyme, whereas PH-20 and Hyal-2 are glycosyl phosphatidyl-inositol (GPI)-linked proteins. The earliest known Hyal is PH-20, the testicular Hyal, which is necessary for ovum fertilization. It is also observed in other normal tissues, and it has been identified in malignancies. Hyal-2 degrades HA into 20 kDa oligosaccharides (about 50 disaccharide units), suggesting a significant role of this isoform in many biological events. Hyal-1 is the serum Hyal and it is expressed in several tissues, with the highest expression in liver, followed by kidney, spleen, and heart. It is also expressed in lung and placenta at very low levels. It is a very important enzyme in prostate and bladder cancers and its presence correlates with stage, aggressiveness, and mortality of at least bladder cancer [3]. Hyal-1 has also been purified from human urine, where it is expressed as two molecular forms. Although Hyal-1 has high specific activity for degrading HA, its concentration in human serum is low (60 ng/mL) [13]. Many different assays regarding the detection and quantification of Hyals have been proposed so far. They are generally based on the answer required. More specifically, the questions raised are concerning either the nature of the enzyme or its total activity. Two different approaches have been proposed so far, namely zymography and analysis of the degradation products. Additional techniques have also been proposed and are applied, such as western blotting and ELISA detection and quantification. However, both latter methods have disadvantages, since they detect mainly the protein nature of Hyals and not their activity. Thus, they cannot distinguish between active and inactive enzymes, i.e., lacking part of the catalytic site. Zymography is a technique used for the detection of active enzyme(s) present in either crude or purified samples. Its advantages are that the enzymatic band(s) is(are) separated from any inhibitor in the enzyme preparation, protein contaminants are well separated from the enzymatic band(s), and different enzymatic bands are well separated and thus the presence of different isoforms may be identified. The method requires the preparation of a polyacrylamide gel into which the substrate—HA—is entrapped. The samples are prepared and subjected to electrophoresis either in neutral or in

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sodium dodecyl sulfate (SDS) conditions. After electrophoresis, SDS is removed through washings in a neutral detergent solution, the gel is incubated overnight in a suitable buffer, and the next day the non-degraded HA is stained with alcian blue. The HA-degrading enzymatic bands appear as clear bands in a blue background. When crude enzymatic preparations are examined for the presence of any Hyal(s), it is better to include an incubation step with pronase to degrade any proteins that may interfere with Hyal detection or to stain with Coomassie, after alcian staining, to detect extra proteins in the gel. With a slight modification, the assay can also be used to detect the presence of any Hyal inhibitors of protein nature. Zymography is followed by an assay through which quantification of Hyal activity is achieved. A variety of different assays have been proposed and all of them involve mixing of the enzyme preparation with hyaluronan and incubation for 4–6 h at 37
C in a suitable buffer. According to the details of the method used, quantification of enzymatic units may be achieved either directly by the Morgan and Elson reaction [14, 15] or after the separation of the obtained products by HPLC [16] or FACE [17]. In the latter, derivatization of the products is a prerequisite, whereas derivatization in HPLC is not obligatory. Moreover, for direct determination of Hyal activity, various plate assays have been proposed, which differ in the type of substrate used, i.e., natural or biotin labeled. Since plate assays are superior to other assays in labor and time cost, only this type of assays will be described. These assays involve immobilization of HA on the plate wells, incubation with the biological sample(s) and degradation when Hyal is present and active, and finally measurement of the remaining hyaluronan in the plate wells. By including different concentrations of standard Hyal, a reference curve is obtained which is used to quantify the enzymatic units of Hyal present in the biological samples. The older assays were of limited sensitivity since they used turbidimetry for HA measurement [18], whereas the newer ones involve immobilization of biotinylated hyaluronan to increase sensitivity through the use of avidin–peroxidase and chromogenic or fluorogenic substrates of peroxidase for the analysis [19]. ELISA methods are common in biochemical laboratories because of their many advantages, such as small quantities of samples, analysis of multiple samples on the same time, existence of automation, and good mathematical interpretation of results obtained, among others. These assays tend to be applied in constantly increased number of biochemical methods, without the need for existence of antibodies for the final measurements. Thus, several ELISA-like or microtiter-based assays have been developed. Such an assay for measurement of Hyal activity is described here that does not require specialized biological reagents.

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The procedure starts with the substitution of a part of the free carboxyl groups of HA with biotin, followed by the covalent immobilization of biotinylated HA onto microplate wells. Then Hyal is added to the wells to cleave HA, and at the completion of enzymatic reaction the residual substrate is detected with avidin–peroxidase system. The result of this reaction can be read in an ELISA plate reader. Using serial dilutions of any Hyal standard, a standard curve of Hyal activity can be obtained that can be used to determine Hyal activity from whatever biological sample. With a slight modification, the assay can also be used to detect the presence of any type of Hyal inhibitors.

2

Materials Prepare all solutions with ultrapure water. Usually double-distilled water or deionized water of a sensitivity less than 7 MΩ-cm at 25
C is acceptable. The reagents should be of analytical grade unless there are no commercially available ones. Prepare all reagents at room temperature and store them at temperature indicated. Do not add any preservative (i.e., sodium azide) in buffers. Glass labware must be carefully cleaned and washed with 70% ethanol. Follow all waste regulations for disposing waste materials.

2.1 Electrophoresis/ Zymography Buffers and Solutions

1. Resolving gel buffer (4): 1.5 M Tris–HCl, pH 8.8. Add about 100 mL water to a 1 L glass beaker. Weigh 181.71 g Tris base and transfer to the beaker. Add water to a volume of about 900 mL. Mix and adjust pH with concentrated HCl. Transfer the solution to a graduated cylinder and make up to 1 L with water. The buffer can be stored at 4
C for up to 1 month. 2. Stacking gel buffer (4): 0.5 M Tris–HCl, pH 6.8. Weigh 60.57 g Tris base and prepare a 1 L solution as above. The buffer can be stored at 4
C for up to 1 month. 3. Acrylamide/bis-acrylamide stock solution (T: 30%, C: 2.7%): Weigh 29.2 g of acrylamide monomer and 0.8 g bis-acrylamide (cross-linker) and transfer to a 100 mL glass beaker containing about 80 mL of water. Mix to dissolve, transfer to a graduated cylinder, and make up to 100 mL with water. Then, filter through a 0.45 μm filter. Care should be taken during weighing, transferring of the solutions, and filtering, since acrylamide is neurotoxic, and thus no personnel should be exposed to it. Store at 4
C for up to 1 month, in a dark bottle or in a bottle wrapped with aluminum foil, since acrylamide is hydrolyzed to acrylic acid and ammonia. 4. Ammonium persulfate: Prepare a 10% (w/v) solution in water. Make fresh daily.

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5. N,N,N,N0 -Tetramethylethylenediamine (TEMED): Use the reagent as supplied. Store either at room temperature or at 4
C. In the latter case its pungent smell is decreased. 6. SDS stock solution: 10% (w/v) solution in water. Make fresh every month. Store at room temperature. 7. SDS-PAGE running buffer (see Note 1): 0.025 M Tris–HCl, pH 8.3, 0.192 M glycine, 0.1% SDS. Prepare routinely 10 stock buffer (0.25 M Tris–HCl, 1.92 M glycine). Weigh 30.285 g Tris base and 144,134 g glycine, mix in a glass beaker containing 800 mL water, dissolve, transfer to a graduated cylinder, and make up to 1 L with water. There is no need to adjust the pH. For use, dilute 100 mL of 10 stock buffer with 890 mL water and add 10 mL of 10% (w/v) SDS. The SDS solution is added last, since it makes bubbles. 8. Bromophenol blue, BPB: To prepare a 0.5% (w/v) solution, dissolve 0.5 g BPB in 100 mL water. 9. Isobutanol saturated with water. 10. Sample treatment buffer (2) (see Notes 1 and 2): Mix 2.5 mL of stacking gel buffer, 4 mL of 10% SDS, 0.5 mL 0.5% BPB, 2 mL pure glycerol, and 1 mL water. The solution is stored in aliquots at 20
C. Since SDS precipitates at low temperatures, the buffer needs to be warmed prior to use. 11. Stock hyaluronan solution: Dissolve 100 mg of high-molecular-mass sodium hyaluronan (usually from rooster comb) in 10 mL water by overnight gentle shaking in a roller at 4
C. The solution is stored in aliquots at 20
C. 12. Washing solution: Triton X-100, 3% (v/v). Mix 3 mL of Triton X-100 with 97 mL water. 13. Incubation buffer (see Note 3): 0.15 M NaCl and 0.1 M sodium formate, pH 4.0. Weigh 9 g sodium chloride and 6.8 g sodium formate in a glass beaker, add 900 mL water, dissolve, adjust the pH with 2 M HCl, transfer to a graduated cylinder, and make up to 1 L with water. The solution is stored at 4
C for 2 weeks. This incubation buffer can be used for detection of Hyals of acidic pH optimum and sodium acetate may be used as alternate. When the presence of Hyals of neutral pH optimum is investigated, it is better to replace sodium formate with sodium acetate. 14. Protein-degrading solution: 100 mM Tris–HCl, pH 8.0, 20 mM NaCl, containing 0.1 mg/mL pronase. Weigh 1.211 g Tris base, 117 mg NaCl in a glass beaker, add 90 mL water, dissolve, adjust the pH with 2 M HCl, add 10 mg of pronase (protease from Streptomyces griseus), dissolve, transfer to a graduated cylinder, and make up to 100 mL with water. The solution is stored in aliquots at 20
C.

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2.1.1 StainingDestaining Solutions (Procedure 1)

1. Substrate staining solution A: 0.5% (w/v) alcian blue in 3% (v/v) acetic acid. Prepare 3% acetic acid by adding 30 mL of glacial acetic acid in water to make a solution of 1 L. Weigh 0.5 g of alcian blue dye and dissolve in 100 mL of 3% acetic acid. Store at room temperature. The dye solution is stable for at least 3 months. 2. Protein staining solution: Coomassie brilliant blue (CBB) 0.15% (w/v) in 30% (v/v) methanol and 10% (v/v) acetic acid. Weigh 0.15 g of CBB and dissolve it in 25 mL methanol. Filter through filter paper in a graduated cylinder and make up the volume to 30 mL with methanol. Then add 60 mL water and 10 mL glacial acetic acid. 3. Destaining solution A: 7% acetic acid. Mix 70 mL of glacial acetic acid with water to make up the volume to 1 L. 4. Destaining solution B: 50% methanol and 10% acetic acid. Mix 500 mL of pure methanol and 100 mL of glacial acetic acid with water to make up the volume to 1 L. 5. Destaining solution C: 5% methanol and 7% acetic acid. Mix 50 mL of pure methanol and 70 mL of glacial acetic acid with water to make up the volume to 1 L.

2.1.2 StainingDestaining Solutions (Procedure 2)

1. Fixative/destaining solution: 20% ethanol and 10% acetic acid. Mix 200 mL of pure ethanol and 100 mL of glacial acetic acid with water to make up the volume to 1 L. 2. Substrate staining solution B: 0.5% alcian blue in 20% ethanol and 10% acetic acid. Weigh 0.5 g of alcian blue and dissolve in 100 mL of 20% ethanol and 10% acetic acid.

2.2 ELISA Procedure Reagents and Buffers

1. 2-(N-morpholino)ethanesulfonic acid buffer, 0.1 M, pH 5.0 (MES): Weigh 1.952 g MES in a glass beaker, add 90 mL water, dissolve, transfer to a graduated cylinder, and make up to 100 mL with water. The buffer can be stored for up to 3 months at 4
C. 2. Hyaluronan solution (1 mg/mL): Weigh 100 mg of hyaluronan of high molecular mass and dissolve in MES buffer, under gentle agitation, for at least 24 h at 4
C. The solution must be prepared prior to use. 3. Stock biotin solution, 0.1 M: Weigh 258.3 mg biotin hydrazide and dissolve in dimethyl sulfoxide at a final volume of 10 mL. The solution can be stored for up to 3 months at 4
C. 4. Stock 1-ethyl-3-(3-dimethylaminopropyl) carbodiimide (EDAC) solution, 0.1 M: Weigh 191.7 mg EDAC and dissolve in water at a final volume of 10 mL. The solution must be prepared prior to use.

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5. N-hydroxysulfosuccinimide (Sulfo-NHS) solution: Weigh 36.8 mg of Sulfo-NHS and dissolve in water at a final volume of 100 mL. The solution must be prepared prior to use. 6. Phosphate buffer saline, PBS: 0.154 M NaCl and 0.01 M sodium phosphate, pH 7.4. Weigh 9 g sodium chloride, 0.31 g of NaH2PO4·H2O, and 1.09 g of anhydrous Na2HPO4 in a glass beaker; add 900 mL water; dissolve; transfer to a graduated cylinder; and make up to 1 L with water. The pH of the final solution will be 7.4. The buffer can be stored for up to 2 weeks at 4
C or in aliquots at 20
C for up to several months. 7. Assay buffer A (for lysosomal/acid-active hyaluronidase): 0.1 M formate, pH 3.7, 0.1 M NaCl, 1% Triton X-100, 5 mM saccharolactone. Weigh 9 g sodium formate, 5.844 g NaCl, and 0.961 g saccharolactone in a glass beaker; add 900 mL water; dissolve; adjust the pH with 2 M HCl; add 10 g Triton X-100; mix; transfer to a graduated cylinder; and make up to 1 L with water. The buffer can be stored for up to 1 month at 4
C. 8. Assay buffer B (for neutral-active enzymes): 0.1 M formate, pH 4.5, 0.15 M NaCl, 1% Triton X-100, 5 mM saccharolactone. Weigh 9 g sodium formate, 8.766 g NaCl, and 0.961 g saccharolactone in a glass beaker; add 900 mL water; dissolve; adjust the pH with 2 M HCl; add 10 g Triton X-100; mix; transfer to a graduated cylinder; and make up to 1 L with water. The buffer can be stored for up to 1 month at 4
C. 9. Washing buffer A: PBS containing 2 M NaCl and 50 mM MgSO4. Weigh 116.88 g sodium chloride and 6.018 g magnesium sulfate in a glass beaker, add 900 mL water, dissolve, transfer to a graduated cylinder, and make up to 1 L with water. The buffer can be stored for up to 3 months at 4
C. 10. Washing buffer B: PBS, 2 M NaCl, 50 mM MgSO4, 0.05% Tween 20. Weigh 116.88 g sodium chloride and 6.018 g magnesium sulfate in a glass beaker, add 900 mL water, dissolve, add 0.5 g Tween 20, mix, transfer to a graduated cylinder, and make up to 1 L with water. The buffer can be stored for up to 3 months at 4
C. 11. Citrate-phosphate buffer, 0.1 M, pH 5.3: Prepare a 0.1 M solution of citric acid by weighing 1.921 g of citric acid and dissolving it in water to 100 mL. Prepare a 0.2 M dibasic sodium phosphate solution by weighing 5.365 g of NaH2PO4·7H2O and dissolving it in water to 100 mL. To prepare the citrate-phosphate buffer pH 5.3, mix 22.8 mL of citric acid solution and 26.2 mL of sodium phosphate solution. All solutions are stable for 1 month at 4
C.

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12. Guanidine-HCl, 6 M: Weigh 57.3 g guanidine-HCl in a glass beaker, add water up to 95 mL, dissolve, transfer to a graduated cylinder, and make up to 100 mL with water. The solution can be stored for up to 6 months at 4
C. 13. Hyaluronidase’s standard solutions: Weigh the appropriate amount of commercial reference Hyal to obtain 10 turbidityreducing units (TRU) and gently dissolve to 10 mL of the appropriate assay buffer A or B (see Note 4). The solution can be stored at 20
C in aliquots of 1 mL. Working solutions of Hyal can be prepared daily by serial dilutions of the standard solution, ranging from 1.0 to 1  106 TRU, and kept on ice. Do not vortex the solutions. 14. Avidin biotin peroxidase complex (ABC) solution: The solution must be prepared exactly before use. Follow the instructions of the supplier. Ten mL of the solution is required for the procedure. The buffer usually used is PBS containing 0.1% Tween 20, which is prepared by adding 10 μL of Tween 20 in 10 mL of PBS with gentle shaking. 15. o-Phenylethylenediamine (oPD) solution: The chromogenic substrate solution is prepared exactly before use by dissolving 10 mg oPD (one tablet) in 10 mL of 0.1 M citrate-phosphate buffer, pH 5.3, followed by the addition of 7.5 μL of 30% H2O2. 16. Covalink-NH 96-well plates.

3

Methods

3.1 Detection of Hyaluronidase Activity(ies) Using HyaluronanZymography

1. Preparation of the polyacrylamide gel of desired concentration with HA entrapped to it (see Note 1): Usually gels of 8–10% final concentration of acrylamide are prepared containing HA at a final concentration of 0.15 mg/mL. To do a 10% gel, mix 2.5 mL of resolving gel buffer (4), 3.33 mL of stock acrylamide/bis-acrylamide stock solution, and 3.83 mL water in a 50 mL vacuum conical flask. Degas under vacuum for up to 30 s and then add 0.15 mL of stock hyaluronan solution, 0.1 mL of SDS stock solution, 80 μL of ammonium persulfate, and 10 μL of TEMED and cast gel within a 7.25 cm  10 cm  1.5 mm gel cassette. Allow enough space for stacking gel and gently overlay with isobutanol saturated with water. 2. Preparation of stacking gel (see Note 1): A 4% polyacrylamide gel is prepared by mixing 1.25 mL of stacking gel buffer (4), 0.67 mL of acrylamide/bis-acrylamide stock solution, and 2.94 mL water in a 25 mL vacuum conical flask. Degas under vacuum for up to 30 s and then add 0.1 mL of SDS stock

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solution, 40 μL of ammonium persulfate, and 5 μL of TEMED. Immediately insert carefully a 10-well gel comb to not introduce air bubbles. 3. Preparation of samples for electrophoresis (see Notes 1 and 2): Mix the biological samples (i.e., cell lysate, cell supernatant, serum, plasma, urine, or saliva) with an equal volume of sample treatment buffer (2) and apply directly on gel. When the biological sample is a tissue extract or a crude preparation from cells, it is recommended to heat the samples for 3 min in a boiling water bath. As a positive control for acidic Hyals it is proposed to use human serum diluted (1:20) with water (see Note 4). 4. Electrophoresis run (see Note 1): Electrophoresis takes place at room temperature. Pour electrophoresis tank with running buffer and assemble the electrophoresis apparatus. Electrophorese at a constant voltage of 100 V until the samples enter the resolving gel and thereafter at a constant voltage of 200 V (the current should be 20 mA/gel) until the dye (BPB) reaches 2–3 mm from the bottom of the gel. 5. Removal of SDS from the gel (see Note 1): Once the electrophoresis is completed, take up the gel by the help of a spatula, rinse with water, transfer it to a container with washing solution, and wash the gel for 1 h at room temperature with gentle shaking. Washing may also be performed at 37
C. 6. Preparation for zymography (see Note 3): Discard the washing solution and wash the gel with incubation buffer for 10 min at room temperature with gentle shaking. Washing may also be performed at 37
C. This step may be repeated once more. 7. Zymography (see Notes 3 and 5): Incubate the gel in incubation buffer for 16 h at 37
C with gentle shaking. The following four steps are usually used when purified preparations of Hyals have to be examined (staining-destaining procedure 1). 8. Staining of the HA substrate: Stain the gel with substrate staining solution A for 1 h at room temperature with gentle shaking. 9. Removal of excess alcian blue dye: Destain the gel with destaining solution A for 1 h at room temperature with gentle shaking. Repeat this step once more. 10. Staining of the proteins: Stain the gel with protein staining solution for 30 min at room temperature with gentle shaking. 11. Removal of excess CBB dye: Destain the gel with destaining solution B for 30 min at room temperature with gentle

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shaking, followed by destaining with destaining solution C (two times), each one for 1 h. The previous four last steps 8–11 may be replaced by the following steps, especially when crude biological samples or tissue extracts are examined (staining-destaining procedure 2). 12. Degradation of extra proteins: Transfer the gel into the protein degradation solution and incubate for 2 h at 37
C with gentle shaking. 13. Stabilization of enzymatic bands—Removal of peptides: Fix the gel with fixative/destaining solution for 20 min at room temperature with gentle shaking. 14. Staining of the HA substrate: Stain the gel with substrate staining solution B for 1 h at room temperature with gentle shaking. 15. Removal of excess alcian blue dye: Destain the gel with fixative/destaining solution for 1 h at room temperature. Repeat this step once more. The gel is now ready and can be either photographed and dried for storage or scanned by a digital scanner (Fig. 1) and the digital image can be processed by an image analysis software.

Fig. 1 Zymographic detection of human serum hyaluronidase. Left: Serum proteins were separated in 7.5% PAGE and staining was performed with alcian blue and Coomassie blue sequentially. Right: Serum proteins were separated in 10% PAGE and staining was performed with alcian blue after treating the gels with pronase. Numbers 1–5 refer to different human sera. Hyaluronidase appears as clear band in a blue background

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3.2 Direct Quantitative Measurement of Enzymatic Units by ELISA-Like Method 3.2.1 Preparation of Biotinylated Hyaluronan (b-HA) and its Immobilization onto ELISA Plate Wells

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1. To the appropriate volume of HA solution add Sulfo-NHS solution to a final concentration of 0.184 mg/mL, the appropriate volume of stock biotin solution to obtain a final concentration of 1 mM, and the appropriate volume of stock EDAC solution to obtain a final concentration of 30 μM. Stir overnight at 4
C. Then, remove the excess of biotin and EDAC from the HA-biotin complex formed, by adding guanidineHCl to obtain a concentration of 4 M and dialyzing against 1000 volumes of water with at least three changes. Common dialysis tubing with a molecular weight cutoff of approximately 14,000 is considered suitable for dialysis of HA. The dialyzed, biotinylated hyaluronan (b-HA) can be stored in aliquots at 20
C for up to several months (see Notes 6 and 7). 2. Mix thoroughly Sulfo-NHS solution with the appropriate volume of b-HA and water to obtain final concentrations of 0.184 mg/mL (Sulfo-NHS) and 0.2 mg/mL (b-HA). Pipette the solution into 96-well Covalink-NH plate (50 μL/well). Then, dilute EDAC to 0.64 mM (0.123 mg/mL) in water and pipette 50 μL into the Covalink-NH plate with the b-HA solution. Incubate the plate either overnight at 4
C or for 2 h at 23
C, to achieve the covalent immobilization of b-HA on the microtiter plate wells (see Note 8). Discard the coupling solution and wash the plates three times with washing buffer A. Such modified plates can be stored at 4
C for up to 1 week.

3.2.2 Hyaluronidase Activity Assay

1. Add 100 μL of the appropriate assay buffer A or B (see Note 3) per well to the Covalink-NH plates containing immobilized b-HA and equilibrate. Add, in triplicate, the set of standards and the unknown samples in the appropriate buffer, using 100 μL/well (see Notes 9 and 10). Don’t forget to include positive and negative control wells (no enzyme or no avidinbiotin complex), in triplicate. Incubate samples in plates for 30–60 min at 37
C, depending on the unknown biological samples. 2. Add 200 μL of 6 M guanidine-HCl per well to terminate the enzymatic reaction and wash thrice with washing buffer B, using 300 μL/well. Then add 100 μL/well ABC solution and incubate for 30 min at room temperature (do not add the ABC solution to the negative control wells). 3. Wash the plate five times with washing buffer B (300 μL/well), then add oPD solution (100 μL/well), and incubate the plate in the dark for 5–10 min. Thereafter, read the plate at 492 nm in an ELISA plate reader (Fig. 2) (see Notes 11 and 12).

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Fig. 2 The steps of the ELISA-like procedure for the determination of hyaluronidase activity. As it is clearly shown, hyaluronan is sparingly substituted with biotin, in order to leave enough amount of free carboxyl groups of glucuronate residues to be used for the immobilization of hyaluronan onto the plate wells, as well as to leave enough disaccharide repeats between biotin residues to be free for cleavage by hyaluronidase

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Notes 1. SDS should be omitted in all electrophoresis buffers when HA-degrading activities must be examined in native conditions. 2. Avoid the use of reducing agents (i.e., β-mercaptoethanol or dithiothreitol) in all buffers and especially sample treatment buffer. 3. A variety of buffers should be compared during the overnight incubation step to establish the optimal buffer for a particular system, since in some cases acetate or phosphate buffers affect Hyal activity. Various assay buffers might be compared, to establish the optimal buffer for a particular system. Formate buffers seem to be superior. 4. Zymography allows the detection of Hyal activity in polyacrylamide gels, after electrophoresis in native and SDS-containing buffers, providing information regarding the molecular weight of the enzyme, or optimum pH, when buffers with varying pH are employed. By co-electrophoresing reference Hyal-1, the assay can also provide semiquantitative measurement of Hyal activity of the samples examined. 5. Zymography permits also the detection of Hyal inhibitors of protein nature, if during the overnight incubation step (zymography) Hyal is included [20]. 6. During the last 20–30 years, the interest of biochemists for a sensitive method quantifying Hyal activity has greatly increased. A number of assays have been developed [21–32] and while some of the very recent ones possess specific properties, they do require specialized equipment and reagents. The simplicity and limited time consumption (in comparison to the number of samples analyzed simultaneously) of zymography and microtiter plate assay have never been reached by any of those assays. 7. HA modified under the procedure described in Subheading 3.2.1, step 1, is substituted with approximately one biotin per 90 disaccharide units and provides a strong signal in the avidin–peroxidase system. Furthermore, it leaves considerable number of free carboxylates to proceed with the covalent attachment of b-HA to plate wells and does not interfere with the ability of the Hyal to degrade the modified substrate. 8. Since HA has a pKa of about 3.1, the covalent immobilization of the b-HA to the plate wells is a prerequisite for the application of the ELISA-like method in various acidic pH values to examine for the presence of different enzymes from various sources and quantify their activity.

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9. Usually, the biological samples under investigation for their Hyal activity should be diluted to contain such amounts of enzymatic units to be within the concentration range of the standards. On the other hand, in biological samples of low enzyme quantity, a purification step should be introduced to enrich enzyme activity. It is not recommended, in the latter case, to increase the incubation time. 10. ELISA-like method permits also the detection of Hyal inhibitors of whatever nature in purified biological samples. In such cases, various dilutions of the unknown samples are co-incubated with a defined amount (defined activity) of reference Hyal in either of the assay buffers [19]. Care should be taken when analyzing crude biological preparations, since the co-presence of Hyal and Hyal inhibitors will affect the results. In those cases, serial dilutions of the biological samples have been proposed to be measured, since inhibitors appear to be present in limited amounts than the enzyme. 11. The standard curve in ELISA-like method is generated by the Hyal standards, usually using a semilogarithmic plot. To obtain the Hyal enzymatic units of the unknown biological samples, insert their absorbance values at 492 nm in the standard curve’s equation. 12. Turbidity-reducing unit is a general term describing enzymatic activity. According to the supplier, however, enzymatic activity may be given as “catal” or “NF”, and the standard curve should be created accordingly. References 1. Jacobson A, Rahmanian M, Rubin K, Heldin P (2002) Expression of hyaluronan synthase 2 or hyaluronidase 1 differentially affect the growth rate of transplantable colon carcinoma cell tumors. Int J Cancer 102:212–219 2. Simpson MA, Lokeshwar VB (2008) Hyaluronan and hyaluronidase in genitourinary tumors. Front Biosci 13:5664–5680 3. Morera DS, Hennig MS, Talukder A, Lokeshwar SD, Wang J, Garcia-Roig M, Ortiz N, Yates TJ, Lopez LE, Kallifatidis G, Kramer MW, Jordan AR, Merseburger AS, Manoharan M, Soloway MS, Terris MK, Lokeshwar VB (2017) Hyaluronic acid family in bladder cancer: potential prognostic biomarkers and therapeutic targets. Br J Cancer 117:1507–1517 4. Kiriake A, Madokoro M, Shiomi K (2014) Enzymatic properties and primary structures of hyaluronidases from two species of lionfish

(Pterois antennata and Pterois volitans). Fish Physiol Biochem 40:1043–1053 5. Biner O, Trachsel C, Moser A, Kopp L, Langenegger N, K€ampfer U, von Ballmoos C, Nentwig W, Schu¨rch S, Schaller J, KuhnNentwig L (2015) Isolation, N-glycosylations and Function of a hyaluronidase-like enzyme from the venom of the spider Cupiennius salei. PLoS One 10(12):e0143963 6. Pavan M, Beninatto R, Galesso D, Panfilo S, Vaccaro S, Messina L, Guarise C (2016) A new potential spreading factor: Streptomyces koganeiensis hyaluronidase. A comparative study with bovine testes hyaluronidase and recombinant human hyaluronidase of the HA degradation in ECM. Biochim Biophys Acta 1860:661–668 7. Mo¨ller C, Clark E, Safavi-Hemami H, DeCaprio A, Marı´ F (2017) Isolation and characterization of Conohyal-P1, a hyaluronidase

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quantum dots via a hydrothermal method as a fluorescent probe for hyaluronidase detection. ACS Appl Mater Interfaces 8:11272–11279 30. Fayad S, Nehme´ R, Langmajerova´ M, Ayela B, Colas C, Maunit B, Jacquinet JC, Vibert A, Lopin-Bon C, Zdeneˇk G, Morin P (2017) Hyaluronidase reaction kinetics evaluated by capillary electrophoresis with UV and highresolution mass spectrometry (HRMS) detection. Anal Chim Acta 951:140–150 31. Gao N, Yang W, Nie H, Gong Y, Jing J, Gao L, Zhang X (2017) Turn-on theranostic

fluorescent nanoprobe by electrostatic selfassembly of carbon dots with doxorubicin for targeted cancer cell imaging, in vivo hyaluronidase analysis, and targeted drug delivery. Biosens Bioelectron 96:300–307 32. Tao L, Song F, Xu N, Li D, Linhardt RJ, Zhang Z (2017) New insights into the action of bacterial chondroitinase AC I and hyaluronidase on hyaluronic acid. Carbohydr Polym 158:85–92

Chapter 13 Method for Studying the Physical Effect of Extracellular Matrix on Voltage-Dependent Ion Channel Gating Eleonora Solari and Andrea Moriondo Abstract Divalent cations can change the actual electrical potential at the outer surface of the plasma membrane. They do so by the so-called Gouy-Chapman-Stern effect which is due to the electrical “masking” that certain ions, especially divalents, can exert onto the electrically negative charged polar heads of the membrane phospholipids. Chondroitin sulfates can chelate free calcium ions to a different extent based on the spatial arrangement of their sulfate groups and can thus alter the actual availability of screening divalent ions at the outer membrane surface. Voltage-dependent ion channels sense the actual potential difference between the two sides of the plasma membrane and are thus exquisite and extremely sensitive “devices” able to react to changes in the electrical potential across the membrane. Hence, by recording the shift in the activation curve of well-known voltage-dependent ionic channels it will be possible to study the physical effect of ECM chondroitin sulfates on membrane conductances. Key words Chondroitin sulfates, Charge screening effect, Voltage-gated ionic channels, Gouy-Chapman-Stern, Calcium ions, Chelation

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Introduction Voltage-gated ion channels are integral membrane proteins which can sense the electrical potential difference across the plasma membrane and can open their gates to let ions flow through them. This, in turn, gives rise to an ionic current which can be measured by means of conventional electrophysiological techniques. When the characteristic voltage dependence of the ionic current is already known, these channels can be used as an exquisite and very sensitive tool in order to explore the tiniest variations of the membrane potential [1]. Since the voltage sensor of the ionic channels is buried inside the membrane thickness, it senses the actual electrical potential difference between the intracellular and extracellular sides of the

Davide Vigetti and Achilleas D. Theocharis (eds.), The Extracellular Matrix: Methods and Protocols, Methods in Molecular Biology, vol. 1952, https://doi.org/10.1007/978-1-4939-9133-4_13, © Springer Science+Business Media, LLC, part of Springer Nature 2019

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membrane. One of the parameters that can influence the extracellular membrane charge is the availability of positively charged ions that can interact and neutralize the negatively charged phospholipid heads. Among the possible positively charged ions, the most effective to deploy this “charge screening” effect are divalent cations, calcium being one of the most prominent [2]. This effect, named “Gouy-Chapman-Stern screening” [3], can effectively change the electrical membrane potential of the outer face so that voltage-dependent ion channels respond to it by altering their normal behavior [4]. This change can be easily recorded by simple electrophysiological techniques. Chondroitin sulfates are one of the extracellular macromolecules which are very effective in binding free calcium ions, thus preventing them from screening the negatively charged phospholipid heads of the membrane. Indeed, changes in chondroitin sulfate concentration and/or composition (4-sulfate or 6-sulfate, for instance), in face of the same nominal extracellular calcium concentration, can give rise to a different actual availability of free calcium ions at the very surface of the membrane, thus altering the charge screening effect [5]. Following the Gouy-Chapman-Stern theory, a decrease in extracellular free calcium ions at the outer membrane surface leads to a more negative potential, closer to the intracellular resting one, thus mimicking a depolarized state, while an increase of free extracellular calcium ions at the outer membrane surface leads to a less negative electrical potential, a larger difference with respect to the intracellular one and thus mimicking a hyperpolarized state (Fig. 1). This, in turn, would result in a leftward or rightward shift, respectively, in the current-voltage plot of the calcium current under investigation (Fig. 2). Among the different voltage-dependent ionic channels that can be used to assess this phenomenon, the protocol described in this chapter uses the L-type voltage-dependent calcium current for three main reasons: 1. L-type channels slowly inactivate, so that during ramp protocols it can be reasonable to assume to have a constant number of active channels for at least 100–150 ms. 2. The ionic current is given by Ohm’s equation: I ¼ G ðV m  E i Þ where G is total membrane conductance referred to the ion channels under investigation, Vm is the membrane potential, and Ei is the Nernst equilibrium potential of the ionic species passing through the channels. Given that

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Fig. 1 (a) The actual membrane potential sensed by the voltage-gated ion channel is identical to the bulk Vm when the extracellular free Ca2+ concentration is enough to neutralize all the negatively charged polar head of the outer membrane surface. (b) When the extracellular free Ca2+ concentration is lowered, some of the negative charges are no more neutralized and thus the actual electrical profile close to the outer membrane surface can be substantially different from the bulk one. In the case depicted, notwithstanding the fact that bulk Vm is the same of panel a, the actual electric potential difference across the membrane is much smaller, mimicking depolarization

Fig. 2 Typical I–V plots of calcium current recorded in low or high extracellular Ca2+. ΔI and ΔV are the two main parameters to be quantified in order to study the screening phenomenon

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 2þ  Ca RT  ln  2þ e Ei ¼ zF Ca i Variations in [Ca2+]e modify Ei and thus the peak current value so that its increase or decrease versus the “reference” recording can be used as an internal control that the shifts in the voltage-dependence recorded in the presence of the test solution are genuine and not due to artifacts. 3. Calcium currents usually are not very large, in the hundreds of pA, minimizing the effects of a poor series resistance compensation during recording and potentially not able to induce saturation of the patch-clamp or voltage-clamp amplifier. In this chapter we detail the procedure to obtain electrophysiological recordings from isolated Xenopus laevis rods of the membrane electrical potential shift induced by chondroitin sulfates using the voltage-dependent Ca2+ current as a tool.

2

Materials Prepare all solutions using ultrapure water, with a resistance of 18 MΩ cm at room temperature. Use analytical grade chemicals and soap- and detergent-free glassware. If in doubt, extensively rinse glassware several times with ultrapure water to decontaminate the inner glass surface. Intracellular solution can be prepared once, aliquoted in 1 mL cryovials (sealed vials able to withstand 20  C), and stored at 20  C for 4 weeks at most. Extracellular solutions must be freshly prepared each day of experiment immediately before the experimental session. Use perfectly dry glassware and reuse the same set of glassware for the same solution each time they are prepared, so to minimize possible artifacts due to crosscontamination of chemical traces. At the end of the experiments extensively wash glassware with ultrapure water and let them dry recovered from dust. Do not use dishwashers; do not rinse other glassware with the same set of glass used for these experiments. To weight powdered chemicals do not use scoops or other type of tools but simply and gently use the index finger to tap the vial and pour small amounts of powder in a weighting boat. Only use plastic boats. Since calcium and magnesium chloride salts tend to be very hygroscopic, we prefer to use commercial, analytical grade 1 M stock solutions in water to be diluted at the desired concentration. Materials detailed below assume a patch-clamp experiment on freshly isolated Xenopus laevis rod photoreceptors.

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To record ICa from Xenopus laevis rod photoreceptors, the intracellular solution to be prepared is as follows: 1. Intracellular solution: 95 mM N-methyl-D-glucamine; 25 mM TEA; 2.5 mM MgCl2; 5 mM EGTA; 10 mM HEPES; 4 mM NaCl; pH 7.2; osmolality 238 mOsm/kg adjusted with Dglucose (usually less than 5 mM) (see Notes 1 and 2 for general principles to apply to solutions to be prepared).

2.2 Extracellular Solutions

To record ICa from Xenopus laevis rod photoreceptors, the extracellular solutions to be prepared are as follows: 1. Standard extracellular solution: 110 mM NaCl; 2.5 mM KCl; 1 mM CaCl2; 1.6 mM MgCl2; 10 mM HEPES; pH 7.6, final osmolality 242 mOsm/kg adjusted with D-glucose (usually less than 15 mM) (see Note 3). 2. Extracellular control solution: 60 mM Na-gluconate; 50 mM NaCl; 2.5 mM KCl; 1 mM CaCl2; 1.6 MgCl2; 10 mM HEPES; pH 7.6; osmolality 242 mOsm/kg adjusted with D-glucose (usually less than 15 mM) (see Note 2). 3. Extracellular chondroitin test solutions: 30 mM TEA; 1 mM CaCl2; 2.5 mM KCl; 1.6 mM MgCl2; 10 mM HEPES; 30 mM CS4-2Na (or CS6-2Na); 50 mM NaCl; pH 7.6 with NaOH, osmolality 242 mOsm/kg adjusted with D-glucose (usually less than 7 mM). These solutions were made with chondroitin-4sulfate (CS4) and chondroitin-6-sulfate (CS6), both acquired from Seikagaku (Tokyo, Japan) and weighted in order to obtain a final concentration of 30 mM of each chondroitin sulfate (see Notes 3 and 4).

2.3

Hardware

In order to perform patch-clamp experiments the standard equipment is required, made of: 1. Patch-clamp amplifier (Axopatch from Molecular Devices, EPC7, or similar from list, to cite two of the most common amplifiers used). 2. An acquisition board (Digidata series from Molecular Devices; various National Instruments boards with at least 10 kHz sampling rates and at least 12 bit A/D resolution) (see Note 5). 3. (Optional) Low-pass Bessel filter in series between patch-clamp amplifier and acquisition board. 4. Vibration-damping table (from TMC or other companies). 5. (Optional) Faraday’s cage. 6. Inverted microscope with a 10 and 40 or higher objectives or upright microscope with 10 and long working distance 40 or higher objective.

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Fig. 3 Typical connections and hardware to build a setup for electrophysiology (see Note 7)

7. Coarse and fine mechanical manipulator for the positioning of fast solution changing multipipette tip. 8. Motorized or hydraulic manipulator (several companies sell them, i.e., Scientifica, Eppendorf, Thorlabs, Narishige to cite few) for patch pipette positioning. 9. A multibarreled perfusion head to be positioned as close as possible to the cell being patched, so that the solution flow completely surrounds the cell and creates a liquid extracellular environment different from the surrounding medium (see Note 6). 10. A gravity or pump fed multiperfusion system composed of several syringes and terminating in the multibarreled head close to the cell. 11. Disposable recording chambers (35 mm petri dishes are fine). 12. Pipette puller (Sutter and Narishige are the most common suppliers). 13. (Optional) Microforge for fire-polishing of pipette tips. 14. Computer (PC or Mac) for data acquisition and analysis. All the hardware described above will be connected to form a functional setup for electrophysiology as depicted in Fig. 3 (see Note 7).

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3.1 Pipette Fabrication

1. Pull borosilicate glass pipettes to a pipette tip resistance between 5 and 6 MΩ when measured with standard intracellular and extracellular solutions. Pull at least 10, preferably 20, pipettes and store them covered until filling and use. Discard the unused pipettes at the end of the experiments (see Note 8).

3.2

1. Wash the cells (see Note 9) in the standard extracellular solution three times by gently spinning with a centrifuge and discard the supernatant.

Cell Preparation

2. Resuspend the cell pellet in a volume of standard extracellular solution so that it will be possible to obtain at least ten 20 μL drops of about 100 cells to be plated in the center of each petri dish/recording chamber. 3. Place a 20 μL drop of cell suspension in the center of a petri dish/bottom of the disposable recording chamber. 4. Put dishes at 4  C until recording. Allow at least 30 min in the refrigerator before starting recording to let cells deposit onto the bottom of the dish (see Notes 10 and 11). 5. Prepare a step and a ramp recording protocol for use (see Note 12). 3.3

Patch Recording

Use a different petri dish/recording chamber for each recording, discarding it at the end of the recording. 1. Add 0.5–1 mL of standard extracellular solution to the dish, or a volume that completely covers cells and the whole bottom of the dish (see Note 13). 2. Allow 60 s for the temperature to equilibrate and place the dish onto the stage of the microscope. 3. Fill the pipette with the intracellular solution, put it in the holder onto the amplifier headstage, and then immerse the pipette tip into the recording chamber while continuously applying positive pressure. 4. Find a suitable cell which must not be adherent to its neighbors and possibly as roundish as possible, to minimize space clamp problems. 5. Connect one line of the head with a reservoir containing the same standard extracellular solution in the bath, one solution line for control extracellular solution, and one different perfusion line for each different test solution. 6. Place the perfusion head onto a clean paper towel or separate dish.

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7. Open all the lines simultaneously and set a perfusion flow of not less than 0.5 mL/min. 8. To check continuity of perfusion, close all the lines except the one with the standard extracellular solution. Use this flow to wash the other tips, and then close the last perfusion line. Dry all the tips. 9. Place the multibarreled perfusion tip in the bath, aligned with the cell to be patched. 10. Move the perfusion head so as to mimic the actual solution change movement later in use, to assure a proper functioning of the system during recording. 11. Move the perfusion head slightly away from the cell so that the cell is not aligned with any of the perfusion lines. 12. Approach the cell with the patch pipette and form a gigaseal onto it. Once obtained, set the command voltage to 60 mV (the holding voltage of the stimulation protocols) and go into the whole-cell configuration. Best to have an input resistance of not less than 1.5 GΩ and an access resistance of less than 10 MΩ. 13. Compensate membrane capacitance and series resistance up to slightly below the oscillation limit of the circuit, or at least 70%. This compensation phase is mandatory, especially for the series resistance. 14. Start the perfusion and recording protocol as follows (see Note 14). 3.4

Data Recording

1. Open the standard extracellular solution line. 2. Place the line in front of the cell. 3. Record the step protocol once. 4. Open the control extracellular solution line. 5. Place the line in front of the cell. 6. Close the standard extracellular solution line. 7. Record the step protocol once. 8. Open the first test solution line. 9. Place the line in front of the cell. 10. Close the control extracellular solution line. 11. Record step protocol once. 12. Open extracellular control solution line. 13. Close test solution line. 14. Record step protocol once. 15. Repeat steps 8–14 for each test solution, with a recording in control extracellular solution in between them.

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16. Terminate the series with a recording in control extracellular solution followed by standard extracellular solution. 17. Repeat steps 1–14 (or only use) for ramp protocols if needed (they are faster than step protocols) (see Note 15). 3.5

Data Analysis

1. Analyze data to obtain current-voltage plots of the recorded current. 2. Quantify the shifts in the peak current level, peak current position, and curve foot between control extracellular solution and each test solution, and among the different test solutions (see Note 16).

4

Notes 1. The solution osmolality has been adjusted for Xenopus laevis rod photoreceptors, but for mammal cells an osmolality of 302–310 mOsm/kg has to be attained by increasing D-glucose concentration. Intracellular solution must however be 5–6 mOsm/kg less than the extracellular one, to account for hydraulic pressure buildup during the whole-cell recording. 2. Since the cellular model to be used may contain significant other currents in addition to the calcium current to be recorded, the actual solutions to be used must be formulated in order to block all the contaminating currents. The actual solutions to be used have to include all the blockers for the eventual conductances that are to be avoided to record the current of choice. Drugs can be prepared as 1000 stock solutions (in water or DMSO) and directly added to the final solution at the desired concentration by diluting them. Other chemicals can be used to substitute for particular ionic species. However, among all solutions, the total concentration of chloride ions must be constant. This point is extremely relevant since the actual command voltage given to the cell depends upon the junctional potential between the Ag/AgCl electrode pellet and the solution in which it is immersed, so that any asymmetry in chloride ions gives rise, when solutions are changed during the experiment, in a shift of the command potential that can and will appear in the recorded current plot as a genuine shift due to the screening effect, but it is artifactual and thus to be avoided. As a reference, we reported in Subheading 2 the actual recipes used for recording of L-type calcium current from Xenopus laevis photoreceptors, made by following the aforementioned rules, and by following advice of Notes 1 and 3 for osmolarity and pH.

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3. Standard extracellular solution is used for keeping cells at 4  C, and its pH and osmolality must be changed to 7.4 and 318 mOsm/kg for mammal cells. 4. In order to create control and test solutions it is important to consider the molecular weight of the molecule under study and the properties of the chelating moiety. Given that the putative Ca2+ chelating moiety is supposed to be the disaccharide unit [6, 7], this approach also helps in determining the actual concentration of Ca2+ chelating sites. The example has been made to elucidate that, in order to have a known concentration of Ca2+ chelating sites, given that the disaccharide unit is able to chelate 1 calcium ion, a total concentration of “disaccharide units” had to be present in the test solution. Hence, from MW the actual chondroitin sulfate concentration has been derived. 5. The acquisition board must be capable to sample at a rate of at least 10 kHz while filtering at 2–3 kHz with a low-passing Bessel 8 pole filter to avoid undersampling. 12 bit A/D resolution is essential to capture small currents without using too high gains to risk saturation. 6. This is extremely important for three main reasons: (1) fast solution change is optimal to ensure a short recording period so as not to introduce artifactual drifts; (2) lowest volume of test solutions potentially made with very expensive reagents; and (3) precise control of the extracellular medium surrounding the cell under investigation. Albeit several companies sell “ready to go” multiline perfusion systems with multibarreled heads, we routinely use a gravity-fed system composed of a variable number of reservoirs made with 20–50 mL disposable syringes, PE tubing, electrovalves, and a custom-made multiline perfusion head. The key point in the system is to use separate tubing from the reservoirs to the cell, with no common output. In this way cross-contamination of solution is avoided and results are clearer. Moreover, we try to use the same syringe and tubing for the same solution for the whole lot of experiments, and then replace the whole system for different sets of experiments. 7. The recording chamber that will host the cells to be probed can be easily obtained with a 35 mm petri dish, unless other means of disposable chambers are already in use in the lab. Petri dishes also simplify the preparative steps, because they can be stored at 4  C with the 20 μL drop of cell suspension in the center until use. The multi-reservoir part of the perfusion system can be made following the suggestions of Note 6, and fitted to a multibarreled perfusion head by means of PE tubing of the smallest diameter compatible with a flux of 0.5–1 mL/min.

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The multibarreled head can be obtained by connecting the PE tubing to an array of glass microcapillaries (for instance the ones used for hematocrit determination, 50 μL volume, but also squared ones sold by WPI Inc.). Otherwise, it can be obtained by using a 1 mm OD max PE tubing with its end facing the cell optionally cut at 45 . Try to avoid the manifold type since they have multiple inlets but one single common outlet. The perfusion head must be mounted onto a three-axis mechanical manipulator, even of a “coarse” type, so that it can be placed in proximity of the cell being recorded and translated during the recording. The microscope, upright or inverted, is best positioned onto a vibration-damping table; otherwise it would be impossible to carry on the electrophysiological recording due to vibrations rupturing the gigaseal. Patch-clamp amplifiers come with their own proprietary headstages and can possibly accommodate different third-party electrode holders, which have to be of a correct diameter for the glass capillaries to be used as electrodes. Electrode holder must be for patch clamp, i.e., must have a side port to apply the positive and negative pressures required for seal formation and rupture. Pressure can be applied by mouth or by using a 1 mL syringe without plunger connected to a PE tubing running to the side port of the electrode holder. A little bit of practice is needed to deliver the correct pressure. Otherwise, expensive pressure-controlling devices are present on the market, but to our experience they are not worth the economical effort: mouth is a better, more sensitive means of pressure control. The patch-clamp amplifier is then connected to the optional low-pass Bessel filter by means of 50 Ω coaxial cables. Beware that 75 Ω video coaxial cables are identical to 50 Ω data cables, so care must be taken not to confuse the two. Impedance unbalance can lead to a poor signal-to-noise ratio that should be avoided. Then, the optional filter is connected, with 50 Ω coaxial cables, to the A/D D/A board, and the board is connected by USB or other proprietary cables to the PC/Mac. The output of the board should be connected to the input ports of the patch-clamp amplifier to deliver the correct stimulus to the cell. Usually, board vendors also sell the acquisition programs used to record data and deliver the stimuli to the patch-clamp amplifier, so that companies like LIST or Molecular Devices are a one-stop shop for all the electronic equipment needed (amplifier with headstage and holders, digital board, software). 8. Alternatively, use a 10 mL syringe connected to a freshly pulled pipette, place the shaft of the syringe at the 10 mL mark, immerse the pipette tip in a small vial filled with EtOH 99%,

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and push air into the pipette until the first tiny bubbles emerge from the tip. Record the syringe volume at which this occurs; it should be between 5 and 6 mL marks for a proper pipette tip. Fire-polishing the tip with microforge is optional. We never had to perform this step in order to obtain gigaseals routinely. 9. The aim of this part of the procedure is to obtain, in each petri dish or disposable recording chamber, at least a hundred cells, isolated one from the others, to choose amidst the one to be patched. The path is different starting from primary cells or cell lines, but then the final steps are identical. We employed Xenopus laevis rods because this was the routinely used cellular preparation of our lab, thus a model very well known and whose conductances could easily be blocked by means routinely used. Another option is to transfect cells like HEK or similar with plasmids carrying calcium channels or other channels of choice, and use them for the experiments. However, given the different yield of expression among different transfected batches, it would be advisable to express the ionic current relative to cell membrane area, or to normalize for a naı¨ve current. 10. For photoreceptors, this could also be the case for other cells; a better preservation of cells can be attained if the petri dishes with the 20 μL drop of cells are placed onto an ice bed in the refrigerator at 4  C. 11. Enzymatic digestion can be a problem if too aggressive. Never use trypsin to dissociate cells, or other nonspecific proteases, since they can damage the extracellular loops of the ionic channels and alter the subsequent electrophysiological measures. Use extracellular matrix digestive enzymes such as collagenase and elastase better to resort to mechanical dissociation or mild collagenase digestion. As a rule of thumb, it is better to get a lower yield of better preserved cells than a higher number of potentially damaged cells. 12. Start from a suitable holding potential of 60 mV held for 50 ms, then a step to test potentials from 80 to +80 mV in 10 mV increments lasting 100 ms, and final 50–100 ms at holding potential of 60 mV. For ramp protocols, build a ramp with a 50 ms epoch at 60 mV, then a ramp from 80 mV to +80 mV of 300 ms of duration, and a final epoch at 60 mV of 50–100 ms of duration (Fig. 4). These values are typical for L-type calcium current recording; use longer test steps or ramps if recording slower currents such as If or Ih, and alter the test voltages accordingly. Protocols are made for calcium current, but they have to be altered for recording for instance hyperpolarization-activated currents, like If or Ih, with a holding potential of 40, and steps lasting

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Fig. 4 (a) Typical step protocol for the study of calcium current. (b) Typical ramp protocol for the study of calcium current

500–600 ms from 130 to 40. Avoid ramps with very slow currents because ramp could be too fast for the current to fully develop before the command potential is changed. 13. Do not pour liquid onto the cells at the center, but use a 1 mL pipette to gently add the solution from the side of the dish, to prevent cells from detaching from the bottom and moving around. 14. The key points of the perfusion scheme are to never open or close a line in front of the cell because the sudden change in flow leads to the end of the seal. Open and close lines while the cell is perfused with another line. Follow a perfusion scheme like this: standard extracellular-control-test, and preferably always record in control solution in between test solutions. At the end of the session perform test-control-standard sequence. Avoid using too many test solutions at once. Two might be enough. 15. At the end of the recording discard the dish and the patch pipette. Expect to spend about 60 min for each dish, especially at the beginning of the experiments and when several solutions are used together with step protocols. 16. Ideally, recordings in standard extracellular solution and control extracellular solution are not supposed to be different, or very small differences may arise from the unbalanced Cl concentrations. Instead, shifts are to be expected when comparing the test solutions with their control extracellular solution recording. These shifts witness the different behavior of the extracellular matrix molecules used in the test solutions. Apply statistical tests where appropriate, such as t-tests or ANOVA.

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References 1. Vigetti D, Andrini O, Clerici M et al (2008) Chondroitin sulfates act as extracellular gating modifiers on voltage-dependent ion channels. Cell Physiol Biochem 22:137–146 2. Piccolino M, Vellani V, Rakotobe LA et al (1999) Manipulation of synaptic sign and strength with divalent cations in the vertebrate retina: pushing the limits of tonic, chemical neurotransmission. Eur J Neurosci 11:4134–4138 3. Grahame DC (1947) The electrical double layer and the theory of electrocapillarity. Chem Rev 41:441–501 4. Frankenhaeuser B, Hodgkin AL (1957) The action of calcium on the electrical properties of squid axons. J Physiol 137:218–244

5. Hille B, Woodhull AM, Shapiro BI (1975) Negative surface charge near sodium channels of nerve: divalent ions, monovalent ions, and pH. Philos Trans R Soc Lond Ser B Biol Sci 270:301–318 6. Iozzo RV (1998) Matrix proteoglycans: from molecular design to cellular function. Annu Rev Biochem 67:609–652 7. Hunter GK, Wong KS, Kim JJ (1988) Binding of calcium to glycosaminoglycans: an equilibrium dialysis study. Arch Biochem Biophys 260:161–167

Chapter 14 Methods for Monitoring Matrix-Induced Autophagy Carolyn Chen, Aastha Kapoor, and Renato V. Iozzo Abstract A growing body of research demonstrates modulation of autophagy by a variety of matrix constituents, including decorin, endorepellin, and endostatin. These matrix proteins are both pro-autophagic and antiangiogenic. Here, we detail a series of methods to monitor matrix-induced autophagy and its concurrent effects on angiogenesis. We first discuss cloning and purifying proteoglycan fragment and core proteins in the laboratory and review relevant techniques spanning from cell culture to treatment with these purified proteoglycans in vitro and ex vivo. Further, we cover protocols in monitoring autophagic progression via morphological and microscopic characterization, biochemical western blot analysis, and signaling pathway investigation. Downstream angiogenic effects using in vivo approaches are then discussed using wild-type mice and the GFP-LC3 transgenic mouse model. Finally, we explore matrix-induced mitophagy via monitoring changes in mitochondrial DNA and permeability. Key words Decorin, Endorepellin, LC3, Beclin 1, Mitophagy, LC3-GFP, Starvation, Angiogenesis

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Introduction A vast and complex reservoir, the extracellular matrix (ECM) contributes an abundance of dynamic roles to surrounding cells to maintain tissue homeostasis and sustain development [1–16]. A major constituent of the ECM, proteoglycans fulfill a range of structural and signaling roles that regulate cellular processes including endocytosis [17–20], cell adhesion [21–24], inflammation and wound healing [25, 26], thrombosis [27], angiogenesis [28–31], and autophagy [32–34]. Autophagy is the “self-eating” process by which cells deliver intracellular proteins, lipids, and organelles to lysosomal compartments for degradation and recycling. Independent of nutrient deprivation, several matrix constituents effectively modulate this intracellular degradative pathway, among which are decorin, endorepellin, endostatin, and kringle V [35].

Carolyn Chen and Aastha Kapoor contributed equally to this work. Davide Vigetti and Achilleas D. Theocharis (eds.), The Extracellular Matrix: Methods and Protocols, Methods in Molecular Biology, vol. 1952, https://doi.org/10.1007/978-1-4939-9133-4_14, © Springer Science+Business Media, LLC, part of Springer Nature 2019

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Decorin, a small leucine-rich proteoglycan (SLRP), functions as a pan-receptor tyrosine kinase inhibitor and impedes endothelial cell migration while activating autophagy [32, 36, 37]. Similarly, endostatin is the 20-kDa C terminal fragment of collagen XVIII-α 1, and exerts its angiostatic activity by binding to α5β1 integrin and inducing autophagy through Vps34, Beclin 1, and LC3-II [38, 39]. Kringle V, instead, is an internal proteolytic fragment of plasminogen that induces autophagy in endothelial cells via binding to glucose-regulated protein 78 kDa (GRP78) while exhibiting potent anti-angiogenic effects on endothelial cell migration and proliferation [40]. Endorepellin is the proteolytically processed C-terminal fragment of the substantial heparan sulfate proteoglycan perlecan [41, 42], a proteoglycan with a complex biology [43–48] involved in the regulation of angiogenesis, both developmental and experimental [28, 49–52]. Located in the basement membrane of vascular endothelial cells, endorepellin has recently emerged as a soluble pro-autophagic and anti-angiogenic molecule of the ECM, inhibiting endothelial cell migration, capillary morphogenesis, and proliferation [53–59]. Through its three laminin-like globular (LG) domains and partial agonistic activity, it signals dually through vascular endothelial growth factor receptor 2 (VEGFR2) to induce protracted autophagy and α2β1 integrin for actin cytoskeleton dissolution, leading to an anti-migratory and angiostatic phenotype [55, 60–63]. Downstream of endorepellin signaling through VEGFR2, 50 -adenosine monophosphate-activated protein kinase (AMPK) is phosphorylated at Thr172, leading to inhibition of mammalian target of rapamycin (mTOR) and induction of autophagy markers including paternally expressed gene 3 (Peg3), Beclin 1, LC3-II, and p62 [32, 33, 64, 65] (Fig. 1). These effects mirror that of Torin 1, a selective competitive inhibitor of mTOR [66]. Ultimately, these ECM molecules interact with their respective receptors to elicit downstream signaling pathways that converge on activating players in the core autophagic machinery including Vps34, Beclin 1, and LC3 [32, 33, 64–67]. They represent a persistent, robust link between autophagy induction and angiogenic inhibition controlled by the extracellular microenvironment. A comprehensive study of these matrix proteins requires experimental treatment ranging from in vitro to in vivo models. The most powerful and effective tool for this is to clone and purify these matrix proteins in the laboratory, ensuring the user’s control over the integrity, purity, and quality of the protein. The ability of certain matrix proteins to induce autophagy while inhibiting vascular growth has shown undeniable clinical promise in the identification of therapeutic targets. In the context of tumorigenesis, stimulating autophagy has been shown to be beneficial for preventing cancer development as it limits inflammation, tissue damage, and genome instability [68]. These pro-autophagic matrix

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Fig. 1 Working model depicting the mechanism of action of endorepellin and torin 1 (both red). Endorepellin is an angiostatic and pro-autophagic factor. Torin 1 is a pro-autophagic factor

proteins are showing angiostatic effects in numerous cancer models, including cancer of the lung, breast, esophageal, and prostate [57, 69–73]. This complex interplay between tumor cells and their vascularized stroma has profound effects on cancer growth regulation [3, 74–79] as angiogenic vessels exert paracrine and angiocrine modes of regulation [80]. Here, we measure the effects of matrixmodulated angiogenesis through an ex vivo aortic ring assay. In this assay, the outgrowth of newly formed blood vessels from the mouse

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thoracic aorta is indicative of a successful angiogenic process. The growing vessels recruit smooth muscle cells and pericytes to associate with the endothelial cell tube, de facto recapitulating the anatomy of mature blood vessels in vivo. Furthermore, autophagosomes can be directly observed in transgenic GFP-LC3 mice after autophagy stimulation through starvation. Protocols concerning tissue extraction for biochemical western blot analysis and mouse organ dissection for immunohistochemistry are also comprehensively detailed here. To visualize autophagosomes, electron microscopy remains the best method to clearly depict the double-membrane structure of these vesicles because it allows to precisely distinguish ultrastructures belonging to lysosomes and other vacuoles in the cell. Over the last half-century, scientists have generated novel techniques such as confocal fluorescence microscopy and atomic force microscopy to study the autophagosomes. Immunofluorescence especially comes in handy to analyze different steps of autophagy in which autophagosomal marker proteins such as microtubule-associated light chain β (LC3β) and sequestosome-1 (p62/SQSTM1) are stained or tagged with fluorophores [81]. The levels of expression of these proteins are also detectable by western blotting for further confirmation. Although morphological analysis of autophagosomes via microscopic techniques was the key first step toward the discovery of the phenomenon of autophagy, it was not sufficient to study the functional aspect of the autophagy. In order to identify proteins as autophagic substrates, autophagosome-lysosome fusion is often inhibited via bafilomycin A1 or chloroquine. This impedes degradation of proteins undergoing autophagy and causes a buildup of autophagosomes, both of which can be detected by western blotting or immunofluorescence [82]. Ultimately, this measures autophagic flux, the measure of autophagic degradation activity, and is an effective method to study proteolytic processing of various proteins of interest. Organelle-specific autophagy utilizes distinct molecular pathways for autophagic degradation of the target. As such, unique proteins and factors are often involved for detecting the type of “-phagy” being examined [83]. In the final section, we discuss established methods for evaluating mitophagy (mitochondrial autophagy) [84] that occurs downstream of decorin and endorepellin signaling. Mounting evidence [85] indicates that mitochondrial turnover via specific autophagic processes is a paramount mechanism for mitochondrial quality control and overall cellular homeostasis. Determining the extent of mitochondrial degradation proceeds through two primary means: imaging and biochemistry. Quantitation of mitochondrial specific chaperones residing within the mitochondrial matrix such as Hsp10, Hsp60, and Hsp70; proteases

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such as Lon and AAA proteinases; and outer mitochondrial membrane proteins such as PTEN-induced putative kinase 1 (PINK1), adenine nucleotide translocator (ANT), and voltage-independent anion channel (VDAC1) via immunoblotting and mitochondrial DNA (mtDNA) via qPCR are absolutely essential. Complementing mtDNA analysis is mitochondrial transcription factor A (TFAM) detection via immunoblotting as TFAM coats the mtDNA and is the sole component found in every mtDNA nucleoid [86, 87]. Examining oxidative phosphorylation subunits via immunoblot further reinforce accelerated mitochondrial turnover following stimulation by decorin or endorepellin. Collectively, data gleaned from both mtDNA clearance and matrix or inner mitochondrial membrane proteins are considered as strict criteria for mitophagy. Determining the mitochondrial membrane potential gives a clear assessment of the mitochondrial permeability pore and its sensitivity to increased Ca2+ uptake. Mitochondrial dyes such as JC-1 are useful for measuring this property and is often a harbinger of future mitophagic processes. There is no one singular empirical scientific method or assay to determine, conclusively, the initiation and progression of mitophagy in mammalian cells. However, using a combination of methods, as outlined below, will give a support for this dynamic process. Therefore, we discuss the quantification of the mtDNA and mitochondrial membrane potential determination via JC-1 staining.

2

Materials

2.1 Cloning of Recombinant Proteoglycans and Purification of HisTagged Recombinant Proteoglycans

1. Veriti 96-well thermal cycler. 2. Lysogeny broth (LB) media and LB agar plates (100 μg/mL ampicillin): Prepare 250 mL of LB media dissolving 6.25 g of LB broth powder in 250 mL water. Prepare 500 mL LB agar dissolving 20 g of LB broth with agar powder in 500 mL water. Autoclave liquids. Make 100 mg/mL ampicillin stock: 0.15 g in 1.5 mL water, filter sterilized in epitube, and stored at 20  C. Cool LB broth media and LB agar and add ampicillin (100 μg/mL). Pour agar in 10 cm plates in the presence of a Bunsen burner (~18 mL/plate). Cool and store labeled plates at 4  C upside down to prevent condensation on agar. 3. Competent Escherichia coli cells. 4. Incubating orbital shaker. 5. Glycerol stock (50% glycerol, 50% water). 6. QIAprep Spin Miniprep kit. 7. GeneJET gel extraction kit.

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8. CutSmart buffer, ligase buffer, T4 DNA ligase, NheI and PmeI restriction enzymes. 9. 50 Tris-acetate-EDTA (TAE) buffer (pH 8.5): 242 g Tris base, 57.1 mL acetic acid, 100 mL 0.5 M EDTA; adjust pH to 8.5 and add water up to 1 L. 10. Agarose gel (1%): Microwave 1 or 2 g agarose in 100 mL 1 TAE (2 mL 50 TAE, 98 mL water) for 1 min for 1% and 2% gel, respectively. Wait for 3 min to cool. Add 3 μL ethidium bromide. Pour liquid into gel mold. 11. Nanodrop 2000. 12. Short blades. 13. Human embryonic kidney cells expressing the Epstein-Barr virus nuclear antigen (HEK 293-EBNA cells). 14. OptiMEM (reduced serum with HEPES and sodium bicarbonate). 15. Lipofectamine 2000. 16. Supplemented Dulbecco’s Eagle modified medium (DMEM): 4.5 g/L Glucose, L-glutamine, and sodium pyruvate, 10% fetal bovine serum (heat inactivated and filtered), 1% penicillin/ streptomycin, 225 μg/mL G418 sulfate (with or without 2 μg/mL puromycin). 17. Hanks’ balanced salt solution (HBSS). 18. EDTA-free proteinase inhibitor mini-tablet. 19. Dialysis membrane (MWCO 6–8 kDa, Spectra/Por). 20. PEG 3350/Carbowax. 21. HisPur Ni-NTA resin beads. 22. 1 Equilibration buffer: 20 mM Na3PO4, 300 mM NaCl, and 10 mM imidazole all dissolved in phosphate buffer saline (PBS), pH 7.4. 23. 1 Binding buffer: 25 mM Imidazole in PBS, pH 7.4. 24. 1 Elution buffer: 250 mM Imidazole in PBS, pH 7.4. 25. 10% Polyacrylamide gel: 4.796 mL Water, 2.5 mL acrylamide (39:1 acrylamide/bisacrylamide, 40% solution), 2.5 mL 1.5 M Tris–HCl (pH 8.8), 0.1 mL 10% sodium dodecyl sulfate (SDS), 0.1 mL 10% ammonium persulfate (APS), 4 μL tetramethylethylenediamine (TEMED). 26. Coomassie Brilliant Blue Dye: 450 mL Methanol, 100 mL acetic acid, 450 mL distilled water, 3 g Coomassie Brilliant Blue R250; filtered. 27. Destain solution: 45% Methanol, 10% acetic acid in distilled water.

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1. Attachment factor (0.2% gelatin in PBS). 2. 12-Well cell culture plates. 3. VascuLife EnGS Endothelial Medium (Lifeline Cell Technology) (see Note 1). 4. Human umbilical vein endothelial cells (HUVEC). 5. Sterile 15 mL tubes. 6. Purified recombinant proteoglycan (see Subheading 3.1: Purification of His-tagged Recombinant Proteoglycans). 7. Sterile PBS. 8. Radioimmunoprecipitation assay (RIPA) buffer: 50 mM Tris–HCl (pH 7.5), 150 mM NaCl, 1 mM EGTA, 1 mM EDTA, 1% Triton-X 100, 0.5% sodium deoxycholate, 0.5% SDS, 1 mM sodium orthovanadate, 1 EDTA-free protease inhibitor tablet, 1 μg/mL leupeptin, 1 μg/mL aprotinin, 100 μM TPCK, 1 mM PMSF. 9. Cell scraper. 10. 5 Sample buffer: 25 mL 1.25 M Tris–HCl (pH 6.8), 50 mL glycerol, 10 g SDS, 4 mL 0.6% bromophenol blue.

2.3 Biochemical Analysis of Autophagic Flux and Cell Signaling

1. Antibodies (Table 1). 2. Tris-buffered saline (TBS) 10: For 1 L of solution, 30 g of Tris base, 80 g of sodium chloride (NaCl), and 2 g potassium chloride (KCl) were dissolved in 800 mL water, stirred until the components entered the solution and the final volume was made up to 1 L. 3. Tris-buffered saline tween (TBST) 1: 100 mL of 10 TBS and 1 mL of Tween-20 were dissolved in 900 mL of sterile water. 4. Blocking buffer: 1% Bovine serum albumin (BSA) (w/v) in TBST, store at room temperature. 5. Chemiluminescent substrate: extended duration pico kit.

SuperSignal™

west

6. Transfer cassette unit: Mini trans-blot cell. 7. Visualizing western blots—Image Quant LAS 4000. 8. Torin 1. 9. Chloroquine. 10. Bafilomycin A1. 11. Compound C. 12. SU5416.

dura

IHC dilution 1:1000 1:1000 1:1000 1:10000 1:1000 1:1000 1:1000 1:1000 1:1000 1:1000 IHC Dilution 1:400 1:400 1:400

Primary antibody

Anti-LC3B

Anti-p62/SQSTM1

Anti-ATG9A

Anti-GAPDH

Anti-PINK1

Anti-ANT

Anti-VDAC1

Anti-Hsp60

Anti-Mitofusin 2

Total OxPhos Human Antibody Cocktail

Secondary antibody

Alexa Fluor 488 anti-rabbit IgG

Alexa Fluor 594 anti-rabbit IgG

Alexa Fluor 594 anti-mouse IgG

Table 1 Antibodies for western blot analysis

Rabbit

Donkey

Goat

Source organism

Mouse

Mouse

Rabbit

Rabbit

Rabbit

Rabbit

Rabbit

Rabbit

Rabbit

Rabbit

Source organism

Thermo Fisher

Thermo Fisher

Thermo Fisher

Company

Abcam

Abcam

Cell Signaling

Cell Signaling

Cell Signaling

Cell Signaling

Cell Signaling

Abcam

Abcam

Sigma

Company

Mitochondrial matrix and inner mitochondrial membrane

Outer mitochondrial membrane and endoplasmic reticulum

Mitochondrial matrix

Outer mitochondrial membrane

Outer mitochondrial membrane

Outer mitochondrial membrane

Loading control

Trans-Golgi network and endosome/autophagosome

Autophagosome

Autophagosome

Marker

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2.4 Microscopic Visualization of Autophagy

1. Glutaraldehyde 2% (v/v) in PBS.

2.4.1 Electron Microscopy

4. Embedding medium Spurr’s resin (Spurr’s solution: 33% resin, 66% resin, and 100% resin).

2. Osmium tetroxide 1% (v/v) in PBS. 3. Graded ethanol (40%, 50%, 60%, 70%, 90%, 96%, and 100%).

5. UC7 Ultramicrotome with glass knives and diamond knives. 6. Toluidine Blue O stain. 7. Light microscope. 8. Copper grids. 9. Uranyl acetate (1–5% solution in water). 10. Lead citrate (0.2% of lead citrate in 0.1 N sodium hydroxide). 11. JEOL 100 CX II electron microscope with Advantage HR digital CCD Camera System. 2.4.2 Atomic Force Microscopy (AFM)

1. Dimension icon AFM. 2. Glutaraldehyde 2% (v/v) in Hanks’ balanced salt solution (HBSS): For 10 mL solution 0.2 mL of glutaraldehyde was dissolved in 10 mL HBSS (freshly prepared solution). 3. Desiccation chamber.

2.4.3 Immunofluorescence Microscopy

1. Four-chamber slides (Nunc™ Lab-Tek™ II CC2™ chamber slide system). 2. Glass coverslips (24  60 mm). 3. Fixative solution [4% paraformaldehyde (PFA) in PBS]: For 500 mL solution, 20 g of PFA powder was added to 400 mL of warm 1 PBS solution followed by dropwise addition of 1 N NaOH solution to allow dissolution of PFA, adjust the volume of the solution to 500 mL with 1 PBS, and the pH was rechecked and set to 6.9 with small amounts of hydrochloric acid; working fraction of the solution was stored at 4  C for a month and rest of the solution was aliquoted and frozen at (20  C) for long-term storage (see Note 2). 4. Permeabilization agent: 0.01% Tween-20, 1% BSA in PBS (see Note 3). 5. Blocking agent: 1% BSA in PBS (see Note 4). 6. Antibodies (Table 1). 7. Hard-set mounting medium with DAPI (Vectashield). 8. Confocal microscope: Zeiss LSM780 NLO confocal/multiphoton microscope. 9. Fluorescence microscope: Leica upright fluorescence microscope DM5500B.

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2.5 Monitoring Autophagy and Angiogenesis Ex Vivo 2.5.1 Aortic Ring Assay

1. 3D collagen: Collagen type I rat tail (4.5 mg/mL), 199 media 10 (phenol red free), sterile water, sterile sodium bicarbonate, sterile 1 M sodium hydroxide (see Notes 5 and 6). 2. 50 mL Syringes. 3. 0.22 μm Filters. 4. Wild-type C57BL/6 mice. 5. HBSS. 6. 96-Well plates. 7. 70% (v/v) Ethanol to surface sterilize the mouse body. 8. Scalpel, sharp scissors, dissection forceps, blade, T-pins, paper towels. 9. Biohazard bag. 10. VascuLife EnGS Endothelial Medium. 11. Blocking buffer: 2% BSA in PBS. 12. 4% PFA in PBS. 13. Lectin-1-TRITC. 14. Confocal microscope: Zeiss LSM780 NLO confocal/multiphoton microscope. 15. DAPI diluted 1:1000 in PBS.

2.6 Detecting Autophagy and Angiogenesis In Vivo

1. GFP-LC3 transgenic mice (Riken).

2.6.1 Starvation-Induced Autophagy in GFP-LC3 Mouse Models

5. 500 mL 30% Sucrose in PBS, made fresh each time.

2. 500 mL 4% PFA in PBS filtered. 3. 1 PBS. 4. 500 mL 15% Sucrose in PBS, made fresh each time. 6. Scalpels (1 per mouse group). 7. CO2 chamber and extra cage. 8. Liquid nitrogen in metal bucket. 9. 50 mL Tubes (1 per mouse) filled with ~40 mL 4% PFA in PBS filtered. 10. Flat Styrofoam surface for dissection. 11. 28.5 gauge needles (1 per mouse). 12. Labeled 1.5 mL tubes (labeled with mouse and organ). 13. 1 Tweezer. 14. 1 Dissection scissor. 15. 0.5 M EDTA.

2.6.2 Mouse Tissue Extraction for Western Blot

1. Mortar and pestle. 2. Labeled pairs of tubes for each tissue sample.

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3. Liquid nitrogen in metal bucket. 4. Tissue extraction solution: In 49 mL T-PER (tissue protein extraction reagent, Thermo Fisher) add 500 μL halt protease inhibitor (100) and 500 μL 0.5 M EDTA. 5. Nanodrop 2000. 6. 5 Sample buffer: 25 mL 1.25 M Tris–HCl (pH 6.8), 50 mL glycerol, 10 g SDS, 4 mL 0.6% bromophenol blue. 7. 10% Acrylamide gel: 4.796 mL Water, 2.5 mL acrylamide (39:1 acrylamide/bisacrylamide, 40% solution), 2.5 mL 1.5 M Tris–HCl (pH 8.8), 0.1 mL 10% SDS, 0.1 mL 10% APS, 4 μL TEMED. 8. Microcentrifuge. 2.6.3 Immunohistochemistry of Tissue Angiogenic Markers

1. Optimal cutting temperature (OCT) compound. 2. Tissue mounts (Biopsy Cassettes). 3. Colorfrost Plus Microscope Slides. 4. Glass coverslips. 5. Dry ice. 6. Cryostat machine. 7. Desiccator and vacuum attachment. 8. Blocking solution: 5% BSA in PBS. 9. 1 PBS. 10. Vertical glass slide container. 11. Hard-set mounting medium with DAPI (Vectashield). 12. Antibodies (Table 2).

2.7 Monitoring Mitophagy 2.7.1 Mitochondrial DNA Determination

1. RNAzol B. 2. Extraction buffer (EB): 4 M Guanidine thiocyanate, 50 mM sodium citrate, and 1 M Tris-free base. 3. Polyacryl carrier. 4. Chloroform. 5. Isopropanol. 6. 8 mM NaOH. 7. 1 M Hepes. 8. 100 mM EDTA, pH 8.0. 9. 75% Ethanol. 10. 1 PBS, pH 7.4. 11. 6-Well plastic cell culture plates. 12. PBS. 13. Decorin/endorepellin.

Calbiochem Thermo Fisher

Rabbit Bovine nasal cartilage Griffonia simplicifolia Mouse

1:200 5 μg/mL 1:300 1:200 1:200 IHC Dilution 1:400 1:400 1:400

HA-binding protein, biotinylated

Isolectin GS-IB4 AF594 conjugate

Anti-His6

Anti-LC3B

Secondary antibody

Alexa Fluor 594 anti-rabbit IgG

Streptavidin Alexa Fluor 594

Alexa Fluor 594 anti-mouse IgG

Rabbit

–

Goat

Source organism

Rabbit

Thermo Fisher

Thermo Fisher

Thermo Fisher

Company

Sigma

BD Pharmingen

Abcam

Abcam

Anti-LYVE1

Rabbit

1:50

Company

Anti-CD31

Source organism

IHC dilution

Primary antibody

Table 2 Antibodies for tissue immunohistochemistry

Autophagosome

His6-tagged protein

Endothelial vessels

Hyaluronan

Lymphatics

Endothelial vessels

Marker

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1. Attachment factor (0.2% gelatin in PBS). 2. Four-chamber glass slides. 3. 7.5 mM JC-1 stock solution. 4. Leica upright fluorescence microscope DM5500B.

3

Methods

3.1 Cloning of Recombinant Proteoglycans and Purification of HisTagged Recombinant Proteoglycans 3.1.1 Cloning of Recombinant Proteoglycans and Protein Core Fragments

To monitor matrix-induced autophagy, an optimal experimental strategy involves cloning, synthesizing, and purifying recombinant proteoglycans and protein core fragments in the laboratory. This ensures high purity in the process of protein preparation that is not otherwise available by purchasing proteoglycans from industrial resources. Here, we provide a detailed protocol of the entire process, from cloning the recombinant proteoglycan cDNA into the expression vector to transfecting eukaryotic HEK 293-EBNA cells with the foreign plasmid to purifying the secreted recombinant protein from cell media. In particular, we chose the pCEP-Pu (derivative of pCEP4) expression vector containing a BM40 secretion sequence upstream of the cDNA insertion site [54]. This ensures secretion of the recombinant proteoglycan into the media for effective purification. 1. The full cDNA coding sequence of the proteoglycan region of interest was constructed with a preceding NheI restriction site (GCTAGC) and proceeding with nucleotides encoding for 6 histidine residues followed by the PmeI restriction site (GTTTAAACG) (see Note 7). 2. This cDNA was then cloned into pUC57 (AmpR) (see Note 8). 3. The pUC57 containing the cDNA (pUC57-cDNA) and the empty pCEP-Pu vector were transformed into competent E. coli cells. For this purpose, 25 μL competent cells were mixed with 20 ng DNA and incubated for 30 min on ice. The solution was then incubated at 42  C for 45 s and put back on ice for 2 min. In each transformation, 475 μL warmed LB media (no ampicillin added) was added and placed in a shaker at 37  C for 1 h. 4. A portion (20 μL) of each transformed solution was then aliquoted into separate tubes to make three dilutions in LB media (1:10, 1:100, and 1:000 with negative control). In sterile conditions, each dilution was plated in agar plates containing 100 μg/mL ampicillin and incubated at 37  C overnight (see Note 9). 5. Two colonies were suspended separately in 2 mL broth containing 100 μg/mL ampicillin and placed in 37  C shaking for 6–8 h.

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6. In a separate tube, 500 μL of bacteria culture was suspended with 500 μL glycerol stock. The remaining 1.5 mL was spun down at 5,900  g for 10 min and stored at 20  C for future plasmid purification. 7. Plasmids from cells transformed with pUC57-cDNA and pCEP-Pu were purified from transformed E. coli using the QIAprep Spin Miniprep kit. 8. pUC57-cDNA and pCEP-Pu plasmid underwent double digest with NheI and PmeI. The following was added in a PCR tube: 1 μg plasmid, 5 μg Cutsmart Buffer, 1 μg NheI, 1 μg PmeI, and 42 μg water. The solutions were subjected to the following conditions: 37  C (2–3 h), 65  C (20 min), and stored at 4  C. 9. The uncut and double-digested plasmids were run on a 1% agarose gel with ethidium bromide at 110 V alongside a 1 kb ladder (see Note 10). 10. The linearized pCEP-Pu plasmid and cDNA fragment were cut out of the gel under UV light with short blades and extracted using GeneJET gel extraction kit (Thermo Fisher) (see Note 11). 11. The DNA concentration of the extraction was measured via Nanodrop (see Note 12), and a 10:1 (vector:insert) ratio solution of 75 ng total DNA was made for ligation (see Note 13). 12. Double-digested pCEP-Pu and cDNA were then ligated together. The following was added in a PCR tube: 50 ng cDNA insert, 25 ng pCEP-Pu vector, 2 μL ligase buffer, 1 μL ligase, and water up to 20 μL. The reaction mixture was subjected to the following conditions: 16  C (16 h), 65  C (10 min), and stored at 4  C. 13. Ligated pCEP-Pu-cDNA was transformed into competent E. coli cells using 5 ng ligated plasmid, selected against ampicillin, grown in culture overnight, and purified (see steps 3 and 4) (see Note 14). 14. Using appropriate primers (see Note 15), the cDNA was sequenced and ligated regions of the plasmid to confirm in-frame and correct nucleotide sequence. 15. pCEP-Pu-cDNA was stably transfected into HEK 293-EBNA cells. HEK 293-EBNA cells were grown in 10 cm dish at 37  C in supplemented Dulbecco’s modified Eagle medium (DMEM) until 70–80% confluent (see Note 16). Two tubes were labeled lipofectamine and DNA and the following were added. Lipofectamine tube: 500 μL OptiMEM media, 30 μL lipofectamine 2000. DNA tube: 500 μL OptiMEM media, 10 μg plasmid. 16. Tubes were incubated for 5 min at room temperature.

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17. Contents from the two tubes were mixed together and incubated for 45 min at room temperature. 18. Media from cells were aspirated and 5 mL OptiMEM with DNA-lipofectamine mixture was added gently to cells for 6 h at 37  C (see Note 17). 19. OptiMEM was aspirated and replaced with 10 mL pre-warmed supplemented DMEM overnight. 20. Media was changed again with supplemented DMEM. 21. After 48 h of transfection, add supplemented DMEM with 500 ng/mL puromycin (see Note 18) and grow in puromycin until stable colonies propagate. 22. Maintain stably transfected cell lines in supplemented DMEM with 2 μg/mL puromycin. 3.1.2 Purification of HisTagged Recombinant Proteoglycans

1. HEK 293-EBNA cells transfected with pCEP-Pu-cDNA were propagated until 80–90% confluent (see Note 19). 2. Media from the cells were aspirated and cells were washed with serum-free HBSS. 3. Cells were serum-starved with serum-free supplemented DMEM media with 2 μg/mL puromycin for 72 h at 37  C (see Note 20). 4. Media was filtered in a 0.22 mM filter, and a single EDTA-free protease inhibitor mini-tablet was dissolved in the media by rocking at room temperature (see Note 21). 5. Filtered media was transferred to a Spectra/Por 2 Dialysis membrane and concentrated with PEG/Carbowax to ~30 mL (see Note 22). 6. Media was dialyzed in 2 L sterile water with 0.1 mM phenylmethylsulfonyl fluoride (PMSF) spinning at 4  C for 36 h (see Note 23). 7. Media was dialyzed in 2 L binding buffer with 0.1 mM PMSF for 36 h (see Note 23). 8. An aliquot of 500 μL HisPur Ni-NTA resin beads were pipetted to an open column (see Note 24). 9. An aliquot of 2 mL equilibration buffer and then 2 mL binding buffer were run through the column (see Note 25). Flowthrough was discarded. 10. The dialysate was run through the column 1–3 times, and the unbound flow-through was saved. 11. An aliquot of 4 mL binding buffer was run through the column. 12. An aliquot of 10 mL elution buffer was run through the column and 20 fractions of 500 μL were numbered and collected (see Note 26).

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Fig. 2 Example of purified recombinant endorepellin (ER) run alongside a BSA concentration curve. The 10% acrylamide gel was run at 18 mA, stained with Coomassie Brilliant Blue dye for 30 min, and destained for 36 h. The calculated concentration of this endorepellin purification is 0.25 mg/mL

13. A small aliquot of 2–5 μL per fraction was run on a 10% polyacrylamide gel, stained with Coomassie dye for 20–30 min, and then washed in destain solution for 36 h. 14. Fractions containing recombinant protein were pooled together. The concentration of the purified protein was measured by running electrophoresis with 1 and 2 μL of eluted protein with BSA standards (0.25, 0.5, 1, and 2 μg) (Fig. 2). 15. Optional: To further concentrate final concentration of purified protein, PEG was used for 10–15 min. 3.2 Endothelial Cell Culture, Proteoglycan Treatment, and Preparation of Lysate

1. Approximately 300 μL 0.2% gelatin was incubated in 12-well plates (~4 cm2/well) in 37  C for at least 2 h. 2. After excess gelatin was aspirated, gelatinized 12-well plates were placed under UV light in a laminar flow hood for 10–20 min. 3. Each well was seeded with ~5  104 human umbilical vein endothelial cells (HUVEC) in pre-warmed supplemented VascuLife EnGS Endothelial Medium. Pre-warmed media were replenished every 2 days until cells were grown to confluency (see Note 27). 4. After cells had been confluent for 1–2 days (see Note 28), experimental treatments were initiated. In a sterile 15 mL tube, purified recombinant proteoglycan (200 nM) was added to cell media (500 μL/well), and the solution was incubated at 37  C for 30 min.

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5. Media from cells were aspirated, and pre-warmed proteoglycan treatments were added to wells (500 μL/well) and incubated at 37  C for the designated time period (see Note 29). 6. Posttreatment, cell culture plates were placed on ice, and media were aspirated from wells. Using ice-cold 1 PBS, cells were washed once, and RIPA (200 μL/well) was added for 10–15 min (on ice at room temperature, rocking). 7. Cells were manually scraped on ice. Remaining lysates from each well were vortexed in 50 μL 5 sample buffer, boiled for 2 min, and stored in 20  C for western blot analysis (see Note 30). 3.3 Biochemical Analysis of Autophagic Flux and Cell Signaling

Proteoglycans such as decorin protein core and the C-terminal perlecan fragment endorepellin have been shown to activate autophagy through inducing phosphorylation of AMPKα [65, 88]. Further, canonical activation of AMPKα via phosphorylation at Thr172 in the T-loop region occurs physiologically upon nutrient deprivation and inhibits mTOR activity to activate autophagy [89, 90]. 1. Endothelial cells were seeded and cultured appropriately (see Subheading 3.2, steps 1–5). 2. Endothelial cells were treated with the recombinant proteoglycan the following signaling/flux inhibitors: Bafilomycin A1 (500 nM) or chloroquine (20 μM) inhibits autophagosomelysosome fusion. Compound C (1 μM) inhibits activation of AMPKα. Torin 1 (20 nM) selectively inhibits mTOR and SU5416 (30 μM) and VEGFR1/2 tyrosine kinase inhibitor. 3. Cells were lysed and processed for western blot analysis (see Subheading 3.2, steps 6 and 7). 4. After samples were run through an acrylamide gel and transferred onto a nitrocellulose membrane, membranes were blocked in 1% BSA in TBST, rocking for 1 h at RT. 5. Membranes were incubated overnight in primary antibodies, diluted at 1:1000 in 1% BSA in TBST (see Note 31) (see Table 1). 6. Once the primary antibody was removed the following day, membrane was washed three times in TBST (15 min each). 7. Secondary antibody was added, diluted at 1:4000 for 1 h at RT, and washed with TBST thereafter three times (15 min each). 8. Membrane was treated with chemiluminescent substrate for 2 min and visualized using an ImageQuant machine.

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3.4 Microscopic Visualization of Autophagy 3.4.1 Electron Microscopy (EM)

1. Specimens were fixed in 2% glutaraldehyde (GA) in 0.1 M PBS at RT for 4 h. 2. After GA fixation, specimens were fixed in 1% osmium tetroxide (OT) in 0.1 M PBS at 4  C for 2 h. 3. Post-fixation tissue fragments were dehydrated in a gradient of ethanol solutions (40%, 50%, 60%, 70%, 80%, 90%, 96%, and 100%). 4. Tissue fragments were embedded in plastic resin (Spurr’s solution: 33% resin, 66% resin, and 100% resin) and polymerized overnight at 80  C. 5. The fix and embedded tissue fragments were cut into 0.5–1 μm thick sections using UC7 ultramicrotome with glass knife. Sections were then transferred to glass microscope slides, stained with Toluidine Blue O stain, and observed under a light microscope to check the morphology of the tissue. 6. After visually confirming the intactness of morphological integrity of the tissue fragment, sections were further cut into 100–120 nm thick sections using UC7 ultramicrotome using a diamond knife. 7. Sections were collected on copper grid and stained with uranyl acetate and lead citrate. Briefly, a small petri was placed within a big petri making two chambers. A small petri dish was covered with parafilm and the outer zone filled with 10–15 NaOH pellets. Big droplets of uranyl acetate were put onto the parafilm (covering the smaller petri) with number of droplets corresponding to number of copper grids to be stained. The copper grids were upturned with sample facing the drops and left for 15 min to allow uptake of stain. The grids were removed, dried on paper, and sequentially washed three times in distilled water. The same process was repeated for lead citrate in a similar but separate setup. 8. Thick sections of 100 nm were observed using a JEOL electron microscope. Digital images were created using an Advantage HR Digital CCD camera system that was attached to the electron microscope.

3.4.2 Atomic Force Microscopy (AFM)

1. HUVEC was grown to confluency on attachment factorcoated four-chamber slides (see Subheading 3.2, steps 1–5). 2. Cells were washed with ice-cold PBS 1 and fixed in 2% glutaraldehyde diluted in HBSS. 3. Fixed cells were re-washed with PBS 1 and dried in desiccation chamber.

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4. Cell structure was obtained by probing fixed cells in tapping mode using a nano-sized silicon tip (tip radius R ~10 nm, spring constant ~42 N/m, NCHV-A, BrukerNano). 5. To measure the elasticity modulus of cells, they were grown up to confluency and probed without fixing. 6. An optical microscope was used to locate individual cells before performing nano-indentation at 7 μm/s indentation depth rate using a microspherical tip (R  2.5 μm, k  0.1 N/m) in the same media in which cells were grown (Dimension Icon AFM) (see Note 32). 7. The force-depth loading curve obtained from nanoindentation was fit using Hertz model with the finite cell height correction, and the effective elasticity indentation modulus was calculated. The Poisson’s ratio of the cells was set to 0.5. 3.4.3 Immunofluorescence Microscopy

1. 5  105 HUVEC cells were plated on attachment factor-coated four-chamber slide and grown to confluency (see Subheading 3.2, steps 1–5). 2. The cells were washed with 1 PBS twice and fixed using 100 μL of 4% PFA per well while on ice for 20 min. 3. The PFA was aspirated, and cells were washed again with 1 PBS (see Note 33). 4. 100 μL of the permeabilization buffer was added to each well and rocked for 30 s before washing off the buffer. 5. The cells were blocked with 100 μL 1% BSA for 45 min at RT. 6. Primary antibodies were diluted at 1:200 ratio in 1% BSA solution in PBS and added 200 μL per well for 1 h (Table 1). 7. The cells were washed thrice with 1 PBS for 5 min each. 8. Secondary antibodies were diluted at 1:400 in 1% BSA solution in PBS and added 200 μL per well for 1 h (Table 1). 9. The cells were then washed thrice with 1 PBS for 5 min each. 10. PBS was aspirated, and the walls of the slide were removed with a separator. 11. The slide was dried to evaporate excess PBS, and two drops of mounting media containing DAPI was added per well before overlaying slide with coverslip. 12. The corners of the slide were sealed with clear nail polish and allowed to air-dry for 2 h before visualization. 13. The slide was viewed at 63 using Zeiss confocal microscope or Leica fluorescence microscope (Fig. 3).

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Fig. 3 Visualizing matrix-induced autophagosomes in vitro. Representative immunofluorescence images of HUVEC treated with vehicle (a), endorepellin (200 nM, 4 h) (b), and serum-starved (HBSS, 4 h) (c). Cells were dually stained with antibodies against beclin 1 (red) and LC3 (green), with prominent autophagosomes showing colocalization (yellow, arrows). Scale bar ¼ 10 μm

3.5 Monitoring Autophagy and Angiogenesis Ex Vivo 3.5.1 Aortic Ring Assay

Aortae can be taken from wild-type C57BL/6 mice. Using a purified pro-autophagic matrix protein, aortic rings induced with protracted autophagy showed reduced sprouting, indicative of hindered angiogenesis, while control rings grew proficiently (Fig. 4). 3D Collagen Preparation for Embedding Aortic Rings

1. The following were added to a 1.5 mL tube on ice to form the 3D collagen gel mixture: 350 μL of collagen, 320 μL of autoclaved water, 120 μL of 199 media 10, 280 μL of NaHCO3, and 20 μL of NaOH (see Notes 34 and 35). 2. Aliquots of 50 μL from the mixture were added to each well of a 96-well plate (see Note 36) and placed in a 37  C incubator for 30 min to allow polymerization of collagen gel layer 1. Dissecting and Preparing Aortic Rings

4. The heart and lungs of the euthanized mouse were removed to expose the aorta (see Note 37). The aorta will appear as a fattywhite tube which runs down from the neck region to the diaphragm along the spine. 5. Using a pair of forceps, the thoracic end of the aorta was stabilized and the abdominal end was cut. The aorta was detached from the spine by running a scalpel gently up toward the thoracic end of the aorta where it branches into brachiocephalic artery, left carotid, and left subclavian arteries. 6. The anterior end of the aorta was cut to release the dissected aorta (see Note 38).

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Fig. 4 Confocal microscopy depicting the difference in microvessel growth between vehicle (PBS) and recombinant endorepellin-treated rings. (a) Dense, long sprouts (yellow arrows) emanating from control aortic rings treated with vehicle (PBS). Rings were stained with CD31 (red). (b). Collapsed microvessels in aortic rings treated with endorepellin (200 nM). Rings were stained with CD31 (red). Scale bar ¼ 300 μm

7. The aorta was transferred onto a petri dish on the dissection board and immersed in HBSS. 8. Using a dissection microscope, the morphology of the aorta was verified (see Note 39). Visceral fat was removed around the aorta using a sharp blade. 9. The aorta was cut into small, uniform rings ~0.5 mm in width. 3.5.2 Embedding Aortic Rings into Collagen Type I and Allowing Sprouting

1. Aortic rings were placed one at a time in each well of the 96-well plate layered with 3D collagen (see Subheading 3.5.1). 2. After placing the rings, a second layer of 50 μL collagen gel was added and incubated at 37  C for 30 min (see Subheading 3.5.1, step 1). 3. An aliquot of 100 μL HUVEC VascuLife EnGS endothelial medium was added to each well and the 96-well plate was placed back into the incubator. 4. The 96-well plate was monitored every day to visualize sprouting, and media were changed every 3 days. 5. When vessels began sprouting from the aortic rings, the purified recombinant matrix protein was added to the surrounding media at the appropriate functional concentration (e.g., 200 nM endorepellin). As a control, an equal volume of PBS was added to the media of a separate aortic ring.

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3.5.3 Staining Aortic Rings with Lectin and DAPI to Visualize Sprouting

1. The culture media was removed and the aortic rings were washed with HBSS. 2. Rings were fixed in 100 μL of 4% PFA per well for 2 h rocking in RT. 3. The PFA was aspirated and rings were washed with PBS (see Note 40). 4. Rings were blocked with 2% BSA in PBS for 3 h at RT (see Note 41). 5. Rings were incubated for 3 h at RT (dark) in a 1:200 dilution of lectin-1-TRITC in blocking buffer to stain the endothelial cells. 6. Rings were washed three times in PBS for 15 min each. 7. Rings were stained with DAPI (1:1000) for 10 min (dark). 8. Rings were washed three times with PBS for 15 min each (see Note 42).

3.5.4 Visualization of the Rings

1. The 96-well plate was taken to the confocal facility and visualized with 63 oil objective. 2. Zen black software was used to acquire z-stack images. 3. Image J software was used for 3D reconstruction of images acquired with Zen black. Individual slices of z-stack were converted into a stack and then into maximum-intensity projection image.

3.6 Detecting Autophagy and Tumor Angiogenesis In Vivo

1. From the same cage, an equal number of GFP-LC3 mice were randomly separated into two cages, one for each group (fed vs. fasted) (Fig. 5).

3.6.1 Starvation-Induced Autophagy in GFP-LC3 Mouse Models

2. Starting early in the morning (7–8 AM), food sources from each cage were removed for 25 h. Water supplies were maintained to prevent dehydration. 3. Mice were immediately euthanized and dissected for biochemical and IHC analysis (see below).

Mouse Organ Dissection and Biochemistry

1. One at a time, mice were euthanized in a CO2 chamber and cervical dislocation was performed. 2. Blood was extracted as follows. The mouse abdomen, peritoneum, and chest cavity are cut open with beating heart exposed. Using a syringe, blood was extracted from heart and transferred to tube precoated with 0.5 M EDTA overnight. Tubes were centrifuged at 400  g for 10 min at 4  C, and serum (supernatant) was transferred into a fresh tube using glass pipettes. 3. Organs were extracted (heart, lungs, kidney, liver, tumor). Organs were removed and briefly rinsed in PBS. A small piece

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Fig. 5 LC3 puncta formation in heart tissue of fasted GFP-LC3 transgenic mice. (a–b) Representative immunohistochemistry images of heart sections from GFP-LC3 mice that were fed (a) or fasted for 25 h (b), showing increased LC3 puncta in fasted hearts

of each organ was cut and placed in 4% PFA for IHC. Remaining part was placed in a 1.5 mL tube and flash frozen in liquid nitrogen and stored at 80  C for protein extraction (see Subheading 3.6.2). Rock organs in PFA for 4 h at room temperature (RT). Organs were incubated in 15% sucrose at RT, rocking for 4 h. Organs were incubated in 30% sucrose at RT, rocking overnight. 3.6.2 Mouse Tissue Extraction for Western Blot

1. Tubes with frozen tissue were kept in liquid nitrogen throughout extraction. 2. For each tissue sample: Liquid nitrogen was poured into a mortar and pestle to pre-cool. Tissue was crushed with pestle while submerged in liquid nitrogen (see Note 43). Remaining tissue was stored back at 80  C. Crushed tissue was poured into new labeled tube carefully (cover opening of tube partially with finger to prevent tissue from coming out when liquid nitrogen is evaporated). An aliquot of 1 mL tissue extraction solution was added to tube and vortexed thoroughly. Mortar and pestle were wiped down with dry paper towel for next tissue. 3. Samples were vortexed thoroughly again (~30 s) and centrifuged at 9,300  g for 8–10 min at 4  C. 4. Supernatant was transferred into a new labeled tube, and pellet was discarded. 5. Using the Protein A280 program in the Nanodrop, protein concentration of each sample was measured and volumes for 30 μg were calculated.

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6. In separately labeled tubes, 30 μg of each sample was aliquoted with the appropriate amount of 5 sample buffer, adding water as necessary to normalize volumes. 7. Tubes were boiled for 2 min and run on a 1.5 mm 10% acrylamide gel, transferred, and probed with the appropriate antibodies (see Note 44). 8. Remaining samples were stored at 80  C. 3.6.3 Immunohistochemistry of Tissue Angiogenic Markers

1. Organs were transferred from sucrose to a dry paper towel.

Mounting Tissue in OCT

3. OCT was poured carefully onto labeled tissue mount (see Note 45).

2. Small parts of each organ were cut and the rest put back in 4% PFA for long-term storage at 4  C.

4. Organs were placed in OCT, and more OCT was added to submerge organs (see Note 46). 5. Tissue blocks are placed carefully in 20  C to solidify OCT and stored at 20  C (see Note 47). Cutting Tissue Sections onto Slides

1. Approximately ten glass slides were numbered and labeled per tissue mount. 2. Solidified tissue mounts were placed on dry ice for transportation to cryostat. 3. All tissue mounts were incubated in the 20  C cryostat for 5–10 min to equilibrate. 4. The plastic top of the tissue mount was removed. 5. OCT was dispensed on head of the cryostat chuck and the head of the tissue mount was stabilized on the chuck (see Note 48). 6. Chuck was tightened onto the cryostat sectioning interface. 7. Tissue sections were cut at 8–10 μm (see Note 49). 8. Slides were placed in a vacuum-tight desiccator at RT for 1 h to dry, and then stored at 80  C for future IHC.

Staining Slides for IHC

1. Slides were blocked in 5% BSA in PBS for 1 h, RT (see Note 50). 2. Excess liquid was dried off the slide using a paper towel without disrupting the tissue section. 3. Tissue sections were submerged with primary antibody solution (in 5% BSA in PBS) for 1 h in RT (see Note 51) (Table 2). 4. Slides were carefully washed three times in PBS. 5. Tissue sections were submerged with secondary antibody solution (in 5% BSA in PBS) for 1 h in RT (see Note 51) (Table 2). 6. Slides were carefully washed three times in PBS.

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7. DAPI hard mount was applied to each section (1 drop/section), the coverslip was placed over sections, and clear nail polish was applied to the corners. Slides were stored at 4  C in the dark. 3.7 Mitophagy Monitoring 3.7.1 Mitochondrial DNA Determination

Determining mitochondrial membrane polarization (ΔΨm) proceeds via live-cell staining with the JC-1 vital dye. Mechanistically, JC-1 is a lipophilic cationic dye sensitive to voltage fluctuations. JC-1 accumulates in the inner mitochondrial membrane in response to ΔΨm. A low ΔΨm denotes loss of membrane potential; JC-1 is monomeric and exhibits green fluorescence. However, under conditions of health ΔΨm, JC-1 accumulates proportionately to the ΔΨm and forms JC-1 aggregates, shifting the JC-1 emission spectrum toward a red fluorescence [91]. This can be quantified using ImageJ software. 1. Subconfluent (~75–80%) tumor cells, seeded in 6-well dishes, were treated with 200 nM purified decorin at 37  C. 2. The 6-well dish was gently placed on ice and washed once with ice-cold PBS. 3. A volume of 1 mL of RNAzol B was added to each well and pipetted up and down several times to homogenize cells. The homogenate was transferred to a clean 2.5 mL round-bottom centrifuge tube and place on ice. The wells were washed with an additional 1 mL of RNAzol B and transferred to the appropriate 2.5 mL tube. 4. An aliquot of 100 μL chloroform for every 1 mL of RNAzol B was added and the mixture was vortexed at maximum speed for 20 s. The tubes were incubated for 5 min at RT. 5. The tubes were centrifuged (12,000  g) for 15 min at RT. The RNA-containing aqueous top layer was discarded (see Note 52). 6. An aliquot of 500 μL EB for every 1 mL of RNAzol B was added to the remaining interphase and organic phase layers. The tubes were inverted six to seven times and incubated at RT for 10 min. 7. The tubes were centrifuged (3000  g) for 30 min at 4  C and the upper layer containing genomic DNA and mitochondrial DNA was transferred to newly labeled 1 mL tubes. 8. An aliquot of polyacryl carrier (4 μL for every 1 mL of RNAzol B used) was added followed by the addition of isopropanol (400 μL for every 1 mL of RNAzol B used) to precipitate the DNA species. The tubes were inverted six to seven times and incubated at RT for 10 min. 9. Tubes were centrifuged (12,000  g) for 5 min at 4  C. Supernatant was discarded. The remaining pellet was washed with 1 mL 75% ethanol and incubated at RT for 5 min. Tubes were

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inverted six to seven times to dislodge the pellet from the bottom of the tube and centrifuged (12,000  g) for 5 min at 4  C. The supernatant was discarded. This step was repeated four times to ensure proper washing of the DNA pellet. 10. The pellet was air-dried at RT for 5 min to remove residual ethanol (see Note 53). 11. An aliquot of 30 μL 8 mM NaOH was added to dissolve the DNA pellet followed by centrifugation (12,000  g) for 10 min at 4  C to remove any undissolved material. 12. The supernatant (~30 μL) was transferred to a freshly labeled 200 μL tube. An aliquot of 0.5 μL 1 M Hepes solution was added to bring sample pH to 8 followed by 0.5 μL 100 mM EDTA. This was vortexed for 15 s. The isolated genomic DNA/mitochondrial DNA (gDNA/mtDNA) mixture was stored at either 4  C or 20  C. Repeated freeze/thaw cycles should be avoided. 13. Using 10 ng of freshly isolated DNA per reaction, conventional qPCR was conducted using the following primer sequences (see Note 54): Primer sequences for mtDNA detection of a 0.2 kb fragment of the ND1 gene: forward: 50 -CCCATTCGCGTTAT TCTT-30 /reverse: 50 -AAGTTGATCGTAACGGAA-30 . Primer sequences for gDNA detection of a 0.2 kb fragment of the LPL gene: forward: 50 -GGATGGACGGTAAGAGTGATT-30 / reverse: 50 -ATCCCAGGGTAGCAGACAGGT-30 . 3.7.2 Detection of Mitochondrial Membrane Permeability Via JC-1 Staining

1. The desired number of four-chamber glass slides were gelatinized with 200 μL 0.1% gelatin for 3 h at 37  C. Gelatin was aspirated, and plates were dried under UV-A/B for 30 min. 2. Endothelial or tumor cells were grown to confluency and treated with 200 nM decorin or 200 nM endorepellin for desired amount of time at 37  C (see Note 55). 3. Cells were incubated with 7.5 μM JC-1 for 20 min at 37  C, and then gently washed three times in warm PBS. Add 500 mL of warmed cell-type appropriate media. 4. Live cells were imaged using Leica upright fluorescence microscope DM5500B (Fig. 6).

4

Notes 1. Always add 1% penicillin/streptomycin to cell media. 2. Paraformaldehyde powder is very corrosive to skin and should be handled wearing gloves; once it is added to warm 1 PBS it gets volatile and should be worked with a mask on.

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Fig. 6 Visualizing mitochondrial depolarization via JC-1 staining. (a–d) Representative fluorescence micrographs depicting live-cell imaging of HUVEC after incubation with vehicle (a), endorepellin (6 h) (b), CCCP (1 h, 30 μM) (c), or FCCP (10 min, 500 nM) (d). HUVEC was cultured in nutrient-rich media and incubated with JC-1 (20 min, 7.5 μM) to assess mitochondrial membrane potential. Depolarized (green) compared with polarized (red) mitochondria. Scale bar ~50 μm

3. Cells should be directly transferred from the incubator to the AFM machine, and not be allowed to stand out of the incubator for more than 30 min; this compromises with their buffering ability and induces stress in them. 4. BSA powder is temperature sensitive and should be stored at 4  C always. 5. Collagen type I rat tail can be a little turbid but that does not affect its functionality. Keep the collagen cocktail (mixture of all the components like bases, media, and collagen) on ice to avoid rapid polymerization in the Falcon tube itself. Ice helps to allow transfer of collagen gel to 96-well plates while still in its liquid state. 6. 199 media should be phenol red free to allow visualization of pH change (color change from yellow to red) on addition of bases (NaOH and NaHCO3) to the collagen. Rat-tail collagen I is acidic to begin with since it has been extracted using acetic acid; adding bases allows polymerization of the collagen into gel form. Change in pH should be visible to eliminate any errors in collagen polymerization. 7. Verify that cDNA sequence contains no NheI or PmeI restriction sites. Extra nucleotides may be added to ensure that the cDNA sequence will be in-frame once inserted into the pCEPPu vector. 8. Genscript offers gene synthesis of custom cDNA sequences into pUC57 (AmpR) vectors. 9. Place agar plates upside down in the incubator to prevent condensation on the agar. 10. Run the gel longer to obtain greater separation of bands.

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11. Elute DNA from gel with water to obtain a lower salt solution. 12. Aim for a DNA concentration of 5–20 ng/μL and ~2260/280 ratio. 13. Calculating molar ratio of 10:1 (vector: insert) with 75 ng total DNA: (a) For example: vector/insert ¼ 10 kb/2 kb ¼ 5. (b) Multiplication factor to obtain: 10:1 ¼ 10/5 ¼ 2. (c) Vector ¼ 25 ng. (d) Insert ¼ 25  2 ¼ 50 ng. 14. In miniprep extraction, elute with EB buffer. 15. Design sequencing primers with the following criteria: (a) Melting temperature (Tm) range of 50–65  C. (b) Cannot dimerize. (c) Between 18 and 30 bp in length. (d) GC content of 40–60%. 16. Grow HEK 293-EBNA cells in culture for two passages before transfection. 17. Ensure that handling of lipofectamine and HEK 293-EBNA cells is entirely sterile to avoid contamination in transfection. 18. Expect to see a lot of cell death upon puromycin addition. This ensures stable transfection. 19. Increased confluency is recommended for optimal protein secretion. 20. Cells may be serum starved for 3–5 days for optimal protein yield. 21. For long-term storage, store at 80  C. 22. Dialysis membrane with a lower MWCO is recommended for faster concentration (6–8 kDa recommended). 23. Change dialysis solution once after 12–24 h for optimal dialysis. 24. Use a 1 mL pipette with the pipette tip cut slightly to prevent damaging resin beads. 25. Use glass pipettes to transfer all liquid into column to prevent protein adherence. 26. An aliquot of 500 μL is approximately ten drops. 27. Grow cells in 1 mL media per well in 12-well plate. 28. Wait at least 1 day after cells reach confluency before starting the appropriate treatment. This allows time for cells to fully lay down their basement membrane and extracellular matrix.

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29. Generally, recombinant proteoglycan treatments do not extend beyond 24 h due to protein degradation. 30. In preparation for western blot analysis, measuring the protein concentration of each lysate is generally not necessary since all samples came from fully confluent monolayers of endothelial cells. 31. Membrane can also be blocked in BSA overnight and then treated with primary antibody the following day for 3 h, followed by washing and treating with secondary antibody for 1 h. This step can be optimized depending on the antibody company/quality. 32. After fixing the cells, they can be stored at 4  C for 4–5 days until staining. 33. Use glass pipettes to transfer all liquid into column to prevent protein adherence. 34. It is essential that the constituents of the 3D collagen cocktail are added in the sequence mentioned in the methods section, with bases being added at the end, to allow commencement of polymerization after addition of all the components. Bases make the collagen (which is acidic to begin with) into neutral constituency, thus providing suitable conditions to begin polymerization. 35. 3D collagen cocktail constituents should be thoroughly mixed to allow uniform polymerization across the volume and homogenous texture of the gel. 36. 96-Well plate is ideal for aortic ring assay since bigger well-size plates cause diffusion of growth factors, thereby hindering sprouting. 37. Upon cutting the heart blood will spurt out of the vessels filling up the cavity. Use paper towels to soak the blood. 38. Aorta tends to curl up when the second end is cut (anterior end). Overlay the aorta with few drops of HBSS to smoothen out the tube, after placing it on the dissection board. 39. Esophagus of the mouse also runs close to the aorta and may be mistaken for aorta. Aorta appears as hollow white tube with fat around it, while esophagus is pinker in appearance with very small lumen. 40. Rings can be stored in PBS after fixing with PFA for 1 week until they are stained with antibodies/dyes. 41. Permeabilization of aortic rings was skipped since they were only stained with lectin-1-TRITC dye, which recognizes glycoproteins present in the basal membrane of endothelial cells. If the rings are to be probed for any intracellular protein, they

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should be permeabilized with Tween which is added to PBS in which BSA is dissolved in the blocking step. 42. Rings should be visualized within 1–2 days of staining with lectin-1 and DAPI to avoid diffusion of signal and development of contamination. For long-term storage (up to 1 month), sodium azide can be added to the wells in very low quantity. 43. Make sure to work quickly throughout and triple glove to protect hands from liquid nitrogen. 44. Avoid using mouse antibodies to probe mouse tissue to avoid nonspecific signal. 45. Avoid bubbles in OCT that interferes with cutting sections. Dispense OCT conservatively to keep in the boundary of the tissue mount. Label tissue mount appropriately for each organ or mouse type. 46. Keep organs in the same plane when embedding in OCT for easy sectioning. 47. For long-term storage, keep tissue mounts at 80  C. 48. Let OCT cool on chuck for ~30 s before attaching tissue mount for better adhesion. 49. Initially, set section thickness to 20 μm until tissue in OCT is reached, and then readjust to 8 μm for actual tissue section. Use brush to flatten section as it is being cut, then touch roomtemperature slide to OCT section, and place in slide holder in RT. Depending on the surface area of the tissue section, two tissue sections can be placed on each slide. 50. For washing and blocking, use vertical glass chambers filled with solution that can hold multiple slides for better efficiency. 51. For neater immunostaining, use a wax pencil to draw a circular boundary around the tissue section on the slide. 52. Instead of discarding the aqueous RNA phase, RNA can be isolated from this phase to verify target RNA levels. For example, if a protein has been implicated in mitophagy, one can knock down the target protein via RNAi techniques and verify depletion via the target RNA in the same samples being assayed for mtDNA content. 53. An alternative to air-drying for 5 min, a KimWipe (or sterile Q-tip) can be used to remove the residual ethanol. Great care must be taken to avoid adsorption of the pellet to the KimWipe or Q-tip. 54. Primer stock concentration is at 100 μM and utilized at a working concentration of 10 μM. Calculate fold changes according to the comparative ΔΔCT method using the gDNA CT values as calibrator samples. As an alternative to mtDNA

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quantification via the comparative ΔΔCT method, establishing of a standard curve of known gDNA concentrations can be used to calculate absolute copy numbers of the mtDNA per condition. 55. It is strongly encouraged to run a positive control for the JC-1 staining. Known protonophores include carbonyl cyanide-mchlorophenylhydrazine (CCCP) or carbonyl cyanide-p-trifluoromethoxy-phenylhydrazone (FCCP). It is recommended to use CCCP at 30 μM and FCCP at 500 nM for either 1 h or 15 min, respectively.

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25. Jung M, Lord MS, Cheng B, Lyons JG, Alkhouri H, Hughes JM, McCarthy SJ, Iozzo RV, Whitelock JM (2013) Mast cells produce novel shorter forms of perlecan that contain functional endorepellin: a role in angiogenesis and wound healing. J Biol Chem 288:3289–3304 26. Walraven M, Hinz B (2018) Therapeutic approaches to control tissue repair and fibrosis: extracellular matrix as a game changer. Matrix Biol 71–72:205–224 27. Nugent MA, Nugent HM, Iozzo RV, Sanchack K, Edelman ER (2000) Perlecan is required to inhibit thrombosis after deep vascular injury and contributes to endothelial cellmediated inhibition of intimal hyperplasia. Proc Natl Acad Sci U S A 97:6722–6727 28. Zoeller JJ, McQuillan A, Whitelock J, Ho S-Y, Iozzo RV (2008) A central function for perlecan in skeletal muscle and cardiovascular development. J Cell Biol 181:381–394 29. Andreuzzi E, Colladel R, Pellicani R, Tarticchio G, Cannizzaro R, Spessotto P, Bussolati B, Brossa A, De Paoli P, Canzonieri V, Iozzo RV, Colombatti A, Mongiat M (2017) The angiostatic molecule Multimerin 2 is processed by MMP-9 to allow sprouting angiogenesis. Matrix Biol 64:40–53 30. Wight TN (2018) A role for proteoglycans in vascular disease. Matrix Biol 71–72:396–420 31. Patel VN, Pineda DL, Hoffman MP (2017) The function of heparan sulfate during branching morphogenesis. Matrix Biol 57–58:311–323 32. Buraschi S, Neill T, Goyal A, Poluzzi C, Smythies J, Owens RT, Schaefer L, Torres A, Iozzo RV (2013) Decorin causes autophagy in endothelial cells via Peg3. Proc Natl Acad Sci U S A 110:E2582–E2591 33. Poluzzi C, Casulli J, Goyal A, Mercer TJ, Neill T, Iozzo RV (2014) Endorepellin evokes autophagy in endothelial cells. J Biol Chem 289:16114–16128 34. Ning L, Xu Z, Furuya N, Nonaka R, Yamada Y, Arikawa-Hirasawa E (2015) Perlecan inhibits autophagy to maintain muscle homeostasis in mouse soleus muscle. Matrix Biol 48:26–35 35. Conde-Knape K (2001) Heparan sulfate proteoglycans in experimental models of diabetes: a role for perlecan in diabetes complications. Diabetes Metab Res Rev 17:412–421 36. Gubbiotti MA, Vallet SD, Ricard-Blum S, Iozzo RV (2016) Decorin interacting network: a comprehensive analysis of decorin-binding partners and their versatile functions. Matrix Biol 55:7–21

Monitoring Matrix-Induced Autophagy 37. Neill T, Schaefer L, Iozzo RV (2012) Decorin, a guardian from the matrix. Am J Pathol 181:380–387 38. Rhodes DR, Yu J, Shanker K, Deshpande N, Varambally R, Ghosh D, Barrette T, Pandey A, Chinnaiyan AM (2004) Large-scale meta-analysis of cancer microarray data identifies common transcriptional profiles of neoplastic transformation and progression. Proc Natl Acad Sci U S A 101:9309–9314 39. Chau Y-P, Lin J-Y, Chen JH-C, Tai M-H (2003) Endostatin induces autophagic cell death in EAhy926 human endothelial cells. Histol Histopathol 18:715–726 40. Nguygen TMB, Subramanian IV, Kelekar A, Ramakrishnan S (2007) Kringle 5 of human plasminogen, an angiogenesis inhibitor, induces both autophagy and apoptotic death in endothelial cells. Blood 109:4793–4802 41. Gubbiotti MA, Neill T, Iozzo RV (2017) A current view of perlecan in physiology and pathology: a mosaic of functions. Matrix Biol 57–58:285–298 42. Gonzalez EM, Reed CC, Bix G, Fu J, Zhang Y, Gopalakrishnan B, Greenspan DS, Iozzo RV (2005) BMP-1/Tolloid-like metalloproteases process endorepellin, the angiostatic C-terminal fragment of perlecan. J Biol Chem 280:7080–7087 43. Cohen IR, Murdoch AD, Naso MF, Marchetti D, Berd D, Iozzo RV (1994) Abnormal expression of perlecan proteoglycan in metastatic melanomas. Cancer Res 54:5771–5774 44. Aviezer D, Iozzo RV, Noonan DM, Yayon A (1997) Suppression of autocrine and paracrine functions of basic fibroblast growth factor by stable expression of perlecan antisense cDNA. Mol Cell Biol 17:1938–1946 45. Cohen IR, Gr€assel S, Murdoch AD, Iozzo RV (1993) Structural characterization of the complete human perlecan gene and its promoter. Proc Natl Acad Sci U S A 90:10404–10408 46. Iozzo RV, Pillarisetti J, Sharma B, Murdoch AD, Danielson KG, Uitto J, Mauviel A (1997) Structural and functional characterization of the human perlecan gene promoter. Transcriptional activation by transforming factor-β via a nuclear factor 1-binding element. J Biol Chem 272:5219–5228 47. Iozzo RV, Zoeller JJ, Nystro¨m A (2009) Basement membrane proteoglycans: modulators par excellence of cancer growth and angiogenesis. Mol Cell 27:503–513 48. Iozzo RV, Cohen I (1993) Altered proteoglycan gene expression and the tumor stroma. Experientia 49:447–455

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triple-negative orthotopic breast carcinoma xenograft model. PLoS One 7:e45559 72. Araki K, Wakabayashi H, Shintani K, Morikawa J, Matsumine A, Kusuzaki K, Sudo A, Uchida A (2009) Decorin suppresses bone metastasis in a breast cancer cell line. Oncology 77:92–99 73. Edwards IJ (2012) Proteoglycans in prostate cancer. Nat Rev Urol 9:196–206 74. Karousou E, Misra S, Ghatak S, Dobra K, Gotte M, Vigetti D, Passi A, Karamanos NK, Skandalis SS (2017) Roles and targeting of the HAS/hyaluronan/CD44 molecular system in cancer. Matrix Biol 59:3–22 75. Bissell MJ, Radisky D (2001) Putting tumors in context. Nat Rev Cancer 1:46–54 76. Bissell MJ, Hines WC (2011) Why don’t we get more cancer? A proposed role of the microenvironment in restraining cancer progression. Nat Med 17:320–329 77. Dvorak HF, Weaver VM, Tlsty TD, Bergers G (2011) Tumor microenvironment and progression. J Surg Oncol 103:468–474 78. Egeblad M, Rasch MG, Weaver VM (2010) Dynamic interplay between the collagen scaffold and tumor evolution. Curr Opin Cell Biol 22:697–706 79. Quail DF, Joyce JA (2013) Microenvironmental regulation of tumor progression and metastasis. Nat Med 19:1423–1437 80. Butler JM, Kobayashi H, Rafii S (2010) Instructive role of the vascular niche in promoting tumour growth and tissue repair by angiocrine factors. Nat Rev Cancer 10:138–146 81. Tanida I, Mimematsu-Ikeguchi N, Ueno T, Kominami E (2005) Lysosomal turnover, but not a cellular level, of endogenous LC3 is a marker of autophagy. Autophagy 1:84–91 82. Klionsky DJ, Abdelmohsen K, Abe A, Abedin MJ, Abeliovich H, Acevedo AA, Adachi H, Adams CM, Adams PD, Adeli K, Adhihetty PJ, Adler SG, Agam G, Agarwal R, Aghi MK, Agnello M, Agostinis P, Aguilar PV, AguirreGhiso J, Airoldi EM, Ait-Si-Ali S, Akematsu T, Akporiaye ET, Al-Rubeai M, Albaiceta GM, Albanese C, Albani D, Albert ML, Aldudo J, Algul H, Alirezaei M, Alloza I, Almasan A, Almonte-Beceril M, Alnemri ES, Alonso C, Altan-Bonnet N, Altieri DC, Alvarez S, Alvarez-Erviti L, Alves S, Amadoro G, Amano A, Amantini C, Ambrosio S, Amelio I, Amer AO, Amessou M, Amon A, An Z, Anania FA, Andersen SU, Andley UP, Andreadi CK, Andrieu-Abadie N, Anel A, Ann DK, Anoopkumar-Dukie S, Antonioli M, Aoki H, Apostolova N, Aquila S, Aquilano K, Araki K,

Monitoring Matrix-Induced Autophagy Arama E, Aranda A, Araya J, Arcaro A, Arias E, Arimoto H, Ariosa AR, Armstrong JL, Arnould T, Arsov I, Asanuma K, Askanas V, Asselin E, Atarashi R, Atherton SS, Atkin JD, Attardi LD, Auberger P, Auburger G, Aurelian L, Autelli R, Avagliano L, Avantaggiati ML, Avrahami L, Awale S, Azad N, Bachetti T, Backer JM, Bae DH, Bae JS, Bae ON, Bae SH, Baehrecke EH, Baek SH, Baghdiguian S, Bagniewska-Zadworna A, Bai H, Bai J, Bai XY, Bailly Y, Balaji KN, Balduini W, Ballabio A, Balzan R, Banerjee R, Banhegyi G, Bao H, Barbeau B, Barrachina MD, Barreiro E, Bartel B, Bartolome A, Bassham DC, Bassi MT, Bast RC Jr, Basu A, Batista MT, Batoko H, Battino M, Bauckman K, Baumgarner BL, Bayer KU, Beale R, Beaulieu JF, Beck GR Jr, Becker C, Beckham JD, Bedard PA, Bednarski PJ, Begley TJ, Behl C, Behrends C, Behrens GM, Behrns KE, Bejarano E, Belaid A, Belleudi F, Benard G, Berchem G, Bergamaschi D, Bergami M, Berkhout B, Berliocchi L, Bernard A, Bernard M, Bernassola F, Bertolotti A, Bess AS, Besteiro S, Bettuzzi S, Bhalla S, Bhattacharyya S, Bhutia SK, Biagosch C, Bianchi MW, Biard-Piechaczyk M, Billes V, Bincoletto C, Bingol B, Bird SW, Bitoun M, Bjedov I, Blackstone C, Blanc L, Blanco GA, Blomhoff HK, Boada-Romero E, Bockler S, Boes M, Boesze-Battaglia K, Boise LH, Bolino A, Boman A, Bonaldo P, Bordi M, Bosch J, Botana LM, Botti J, Bou G, Bouche M, Bouchecareilh M, Boucher MJ, Boulton ME, Bouret SG, Boya P, BoyerGuittaut M, Bozhkov PV, Brady N, Braga VM, Brancolini C, Braus GH, Bravo-San Pedro JM, Brennan LA, Bresnick EH, Brest P, Bridges D, Bringer MA, Brini M, Brito GC, Brodin B, Brookes PS, Brown EJ, Brown K, Broxmeyer HE, Bruhat A, Brum PC, Brumell JH, Brunetti-Pierri N, Bryson-Richardson RJ, Buch S, Buchan AM, Budak H, Bulavin DV, Bultman SJ, Bultynck G, Bumbasirevic V, Burelle Y, Burke RE, Burmeister M, Butikofer P, Caberlotto L, Cadwell K, Cahova M, Cai D, Cai J, Cai Q, Calatayud S, Camougrand N, Campanella M, Campbell GR, Campbell M, Campello S, Candau R, Caniggia I, Cantoni L, Cao L, Caplan AB, Caraglia M, Cardinali C, Cardoso SM, Carew JS, Carleton LA, Carlin CR, Carloni S,

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Chapter 15 Method for Determining Gelatinolytic Activity in Tissue: In Situ Gelatin Zymography Elin Hadler-Olsen and Jan-Olof Winberg Abstract To explore the physiological or pathological roles of proteases, it is important to be able to detect and precisely localize them in a tissue, to differentiate between inactive and active forms, as well as to quantify and determine the nature of the enzyme that degrades a given substrate. Here we present an in situ gelatin zymography method that allows for a precise localization of active gelatin-degrading enzymes in a tissue section. In this method, dye-quenched gelatin is put on top of a tissue section. During an incubation period, active gelatinolytic enzymes will degrade the substrate and fluorescent signals are emitted from the locations of these enzymes. Key words In situ gelatin zymography, Tissue, Gelatinase, Proteases, Matrix metalloproteases

1

Introduction Proteases cleave proteins and peptides either at the N- and C-terminal ends (exopeptidases) or within the polypeptide chain (endopeptidases). These enzymes exist in all living organisms and it is estimated that there are more than 66,000 different proteases [1, 2]. They are either localized within a cell, on or in the cell membrane, or secreted from the cell into the extracellular space [3]. Proteases are important for an organism’s survival [4, 5]. Dysregulation of one or several proteases in humans and other vertebrates is associated with disease, and proteases are often involved when microorganisms invade a host [6–9]. Hence, proteases are important targets for therapeutic intervention [10, 11]. Proteases are classified into eight different classes/clans based on the amino acid or prosthetic group involved in the catalytic reaction. These classes are aspartic (A), cysteine (C), glutamic (G), metallo (M), asparagine (N), mixed (P), serine (S), and threonine (T) proteases (Merops database) [12]. There are more than 566 human and 644 murine proteases, of which 273/341 are secreted, 277/283 are intracellular, and 16/16

Davide Vigetti and Achilleas D. Theocharis (eds.), The Extracellular Matrix: Methods and Protocols, Methods in Molecular Biology, vol. 1952, https://doi.org/10.1007/978-1-4939-9133-4_15, © Springer Science+Business Media, LLC, part of Springer Nature 2019

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are intramembraneous, respectively. The majority of human and murine proteases are of the metallo, serine, and cysteine types, where most of the metallo- and serine proteases are secreted while the cysteine proteases are mainly localized within the cell [3]. Proteases induce an irreversible change of the substrate they process. Hence, most proteases are tightly regulated at the transcriptional, posttranscriptional, translational, and/or the posttranslational level. These enzymes are synthesized in an inactive pro-form, and are activated either within the cell or in the extracellular space. Once activated, their activity is regulated by protease inhibitors that bind either reversibly or irreversibly to the enzyme [6, 9, 13, 14]. To explore the physiological or pathological roles of proteases, it is important to be able to detect and precisely localize them in a tissue, to differentiate between inactive and active forms, as well as to quantify and determine the nature of the enzyme that degrades a given substrate. Here, we present a protocol for the detection of active gelatinases in a tissue, i.e., in situ gelatin zymography. There are several enzymes that can degrade gelatin (denatured collagen), and two of them are the two matrix metalloproteases MMP-2 and MMP-9 [15]. Serine proteases like trypsin, plasmin, and matriptase as well as cysteine proteases such as cathepsin L can also degrade gelatin [16–19]. In this protocol we focus on proteases that function around a neutral pH. In situ gelatin zymography allows for a precise localization of active gelatin-degrading enzymes in a tissue section. Dye-quenched gelatin is put on top of a tissue section, and during an incubation period active gelatinolytic enzymes will degrade the substrate and fluorescent signals are emitted from the locations of these enzymes. Originally, in situ zymography was only performed on frozen tissue sections because formalin, the most commonly used fixative for histology/histopathology practice, disrupts proteolytic activity. Compared to fixed, paraffin-embedded tissue sections, frozen sections are generally thicker and the tissue morphology is more difficult to interpret. This can hamper the precise localization of the enzyme activity by in situ zymography. We have demonstrated that some fixatives, including ethanol and a zinc buffer-based fixative (see Note 1), preserve the activity of gelatinolytic enzymes, allowing in situ zymographic analyses of fixed, paraffin-embedded tissue with superior morphology [20]. Immunofluorescence is a frequently used method to localize specific proteases in a tissue and can be used along with in situ zymography to establish if a specific protease co-localizes with protease activity detected by in situ zymography. Another option is to extract proteases from the tissue used for in situ zymography, and analyze the extracted proteins by gelatin SDS-PAGE zymography (see protocol for Chapter 16 by Hadler-Olsen and Winberg in this volume). For both in situ and SDS-PAGE gelatin zymography, various types of inhibitors can be

In Situ Gelatin Zymography

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Fig. 1 In situ zymography of zinc buffer-fixed mouse mammary gland tissue. Gelatinolytic activity is seen as green fluorescence in ducts (a) due to cleavage of DQ-gelatin in the working solution. Addition of 20 mM EDTA to the working solution (b) inhibits all gelatinolytic activity, indicating that metalloproteases are responsible for the gelatinolytic activity. Nuclei are stained blue by DAPI

used to determine the type of protease responsible for the substrate degradation (see Note 2). Figure 1 shows an example of our in situ zymography method, using zinc buffer-fixed mouse mammary gland tissue.

2

Materials

2.1 Reagents and Buffers

1. Xylene or Histoclear (for paraffin-embedded tissue). 2. 100%, 95%, and 70% ethanol (for paraffin-embedded tissue). 3. Adhesive/positively charged glass slides. 4. Formalin 10% (for frozen sections). 5. Fluorescent mounting media, coverslips. 6. Optional: Enzyme inhibitors such as EDTA, Galardin, Pefabloc. 7. Optional: Nuclear stain such as DAPI or propidium iodide (PI). DAPI solution: DAPI stock (10 mg/mL) is diluted 1:10,000 in PBS. 8. Phosphate-buffered saline (PBS): 10 mM Phosphate, 2.7 mM KCl, and 137 mM NaCl, pH 7.4. 9. In situ zymography reaction buffer: 50 mM Tris–HCl, pH 7.6, 150 mM NaCl, 5 mM CaCl2, and 0.2 mM sodium azide.

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10. Dye-quenched (DQ) gelatin: Dissolve 1 mg DQ gelatin (1 vial) in 1.0 mL Milli-Q water. Protect the substrate from light. The dissolved substrate can be stored in the dark at 4  C for some time (at least 2 months). 11. DQ gelatin working solution and control solutions: Dilute the dissolved substrate (DQ gelatin) 1:50 in the in situ zymography reaction buffer to make the working solution for in situ zymography. Calculate 250 μL of working solution per section. To evaluate the contribution of various enzyme classes to the gelatinolytic activity, working solutions containing both DQ gelatin and enzyme inhibitors can be used on control slides. See Note 2 regarding controls. To control for autofluorescence in the tissue, the in situ zymography reaction buffer without substrate (DQ gelatin) added can be used on control slides. 2.2

Equipment

1. Microtome (for paraffin-embedded tissue) or cryostat (for frozen tissue). 2. Heating cabinet (37  C and 58  C). 3. Humidity chamber (opaque). 4. Fluorescent microscope or confocal microscope with filters/ lasers to detect FITC and DAPI (optional).

3

Methods

3.1 Cut Tissue Sections 3.1.1 Zinc Buffer-Fixed or Ethanol-Fixed, ParaffinEmbedded Tissue

3.1.2 Frozen Tissue

1. Cut 4–5 μm thick tissue sections on a microtome. 2. Put a tissue section on an adhesive/positively charged glass slide. 3. Incubate in a heating cabinet set to 58  C for about 3 h to let the section adhere to the slide and to remove most of the paraffin. 1. Cut about 10 μm thick sections. 2. Put a tissue section on an adhesive/positively charged glass slide. 3. Let the tissue adhere to the glass slide by incubating it at room temperature for about 30 min.

3.2 Preparation of Tissue Sections 3.2.1 Deparaffinization and Rehydration of Fixed, Paraffin-Embedded Sections

1. Deparaffinize the sections in xylene or histoclear (two baths, 10 min each). 2. Rehydrate the sections in graded alcohol baths (100% ethanol 5 min  2, 95% ethanol 5 min  2, 70% ethanol 5 min  2, water). 3. After rehydration, it is important to keep the sections humid.

In Situ Gelatin Zymography 3.2.2 Removal of Traces of Embedding Medium Such as OCT from Frozen Tissue

3.3 Incubation of Tissue Sections with DQ Gelatin and Analysis

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1. Rinse the sections in a PBS bath for 5 min. 2. Keep the sections humid during the remaining procedures.

Incubate tissue sections with either DQ gelatin working solution or control solution. Remember to protect the sections from light in all steps below. 1. Pipet 250 μL of the DQ gelatin working solution or control solution (in situ zymography reaction buffer) to each tissue section. 2. Cover with a piece of parafilm, the size of the slide. 3. Incubate in a dark humidity chamber at 37  C. Incubation time must be optimized for the tissue of interest. It is wise to first run a pilot with different incubation times from 30 min to 6 h. 4. Remove the parafilm gently. 5. Rinse off the substrate with Milli-Q water—do not direct the water flow straight on the tissue. 6. Rinse the sections further in PBS baths (5 min  2). 7. If using frozen, unfixed sections: After step 6 above, fix the section by adding 500 μL of 10% formalin to the section or put in a formalin bath for 5 min (in a safety cabinet). Remove the formalin and rinse as above. 8. Optional (for both fixed, paraffin-embedded tissue and frozen tissue): To help orienting the tissue, you can counterstain the nuclei with a fluorescent nuclear stain such as DAPI or PI. Some fluorescent mounting media contain DAPI or you can do a separate DAPI staining prior to mounting. If separate DAPI staining is performed, the following protocol may be used: sections are incubated with DAPI solution for 5 min. Rinse with PBS (5 min  2) and then mount in a mounting medium for fluorescence. 9. Mount a cover glass over the tissue section and seal with nail polish. 10. Analyze in fluorescence microscope or confocal laser microscope. The DQ gelatin is labeled with (fluorescein isothiocyanate) FITC which has a λex at 495 nm and λem at 517 nm. The DAPI has a λex at 358 nm and λem at 461 nm. The fluorescence of FITC is pH dependent (see Note 3).

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Notes 1. Tissue fixation: Formalin disrupts all gelatinolytic activity, whereas some fixatives preserve enzyme activity such as 70% ethanol or zinc buffer fixative. The formula for the zinc buffer fixative is as follows: calcium acetate 0.5 g, zinc chloride 5.0 g, zinc acetate 5.0 g, and dissolve in 1 L 0.1 M Tris–HCl buffer, pH 7.4. Mix to dissolve. The final pH will be approximately 6.8 (6.5–7.0). 2. Control slides: A number of different enzymes can degrade gelatin. To evaluate the contribution of specific enzymes or classes of enzymes, inhibitors may be added to the in situ zymography reaction buffer. Make sure to add a sufficient concentration to inhibit the enzymes of interest. EDTA inhibits all metallo-dependent enzymes, galardin inhibits matrix metalloproteases, pefabloc inhibits serine proteases, etc. Combinations of inhibitors can also be used. 3. The fluorescence of FITC is pH dependent with a plateau between pH 7 and 9. The fluorescence decreases drastically below pH 7 [21] and hence this DQ gelatin is not recommended for studies of gelatin-degrading enzymes with a low pH optimum.

References 1. Artenstein AW, Opal SM (2011) Proprotein convertases in health and disease. N Engl J Med 365:2507–2518 2. Winberg JO (2012) Matrix proteinases: biological significance in health and disease. In: Karamanos NK (ed) Extracellular matrix: pathobiology and signaling. De Gruyter, Berlin, pp 230–238 3. Overall CM, Blobel CP (2007) In search of partners: linking extracellular proteases to substrates. Nat Rev Mol Cell Biol 8:245–257 4. Quiros PM, Langer T, Lopez-Otin C (2015) New roles for mitochondrial proteases in health, ageing and disease. Nat Rev Mol Cell Biol 16:345–359 5. Ricard-Blum S, Vallet SD (2016) Proteases decode the extracellular matrix cryptome. Biochimie 122:300–313 6. Hadler-Olsen E, Fadnes B, Sylte I et al (2011) Regulation of matrix metalloproteinase activity in health and disease. FEBS J 278:28–45 7. Maeda H (1996) Role of microbial proteases in pathogenesis. Microbiol Immunol 40:685–699

8. Tokito A, Jougasaki M (2016) Matrix metalloproteinases in non-neoplastic disorders. Int J Mol Sci 17. https://doi.org/10.3390/ ijms17071178 9. Turk V, Stoka V, Vasiljeva O et al (2012) Cysteine cathepsins: from structure, function and regulation to new frontiers. Biochim Biophys Acta 1824:68–88 10. Brotz-Oesterhelt H, Sass P (2014) Bacterial caseinolytic proteases as novel targets for antibacterial treatment. Int J Med Microbiol 304:23–30 11. Gialeli C, Theocharis AD, Karamanos NK (2011) Roles of matrix metalloproteinases in cancer progression and their pharmacological targeting. FEBS J 278:16–27 12. Rawlings ND, Barrett AJ, Thomas PD et al (2018) The MEROPS database of proteolytic enzymes, their substrates and inhibitors in 2017 and a comparison with peptidases in the PANTHER database. Nucleic Acids Res 46: D624–D632 13. Gaffney J, Solomonov I, Zehorai E et al (2015) Multilevel regulation of matrix metalloproteinases in tissue homeostasis indicates their

In Situ Gelatin Zymography molecular specificity in vivo. Matrix Biol 44–46:191–199 14. Overall CM, Lopez-Otin C (2002) Strategies for MMP inhibition in cancer: innovations for the post-trial era. Nat Rev Cancer 2:657–672 15. Malla N, Sjoli S, Winberg JO et al (2008) Biological and pathobiological functions of gelatinase dimers and complexes. Connect Tissue Res 49:180–184 16. Heussen C, Dowdle EB (1980) Electrophoretic analysis of plasminogen activators in polyacrylamide gels containing sodium dodecyl sulfate and copolymerized substrates. Anal Biochem 102:196–202 17. Lin CY, Anders J, Johnson M et al (1999) Molecular cloning of cDNA for matriptase, a

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matrix-degrading serine protease with trypsinlike activity. J Biol Chem 274:18231–18236 18. Spens E, Haggstrom L (2005) Protease activity in protein-free NS0 myeloma cell cultures. In Vitro Cell Dev Biol Anim 41:330–336 19. Winberg JO, Gedde-Dahl T (1986) Gelatinase expression in generalized epidermolysis bullosa simplex fibroblasts. J Invest Dermatol 87:326–329 20. Hadler-Olsen E, Kanapathippillai P, Berg E et al (2010) Gelatin in situ zymography on fixed, paraffin-embedded tissue: zinc and ethanol fixation preserve enzyme activity. J Histochem Cytochem 58:29–39 21. Geisow MJ (1984) Fluorescein conjugates as indicators of subcellular pH. A critical evaluation. Exp Cell Res 150:29–35

Chapter 16 Method for Determining Gelatinolytic Activity in Tissue Extracts: Real-Time Gelatin Zymography Elin Hadler-Olsen and Jan-Olof Winberg Abstract To explore the physiological or pathological roles of proteases, it is important to be able to detect and precisely localize them in a tissue, to differentiate between inactive and active forms, as well as to quantify and determine the nature of the enzyme that degrades a given substrate. Here we present a protocol for real-time gelatin zymography that is very useful for the detection of gelatin-degrading proteases in tissue extracts. This method uses fluorescence-labeled gelatin and therefore we also present an easy, fast, and cheap method for labeling gelatin with 2-methoxy-2,4-diphenyl-3(2H)-furanone (MDPF). Key words Gelatinase, Fluorescence labeling of gelatin, MDPF-gelatin, Real-time gelatin zymography, Tissue, Proteases, Matrix metalloproteases

1

Introduction Proteases cleave proteins and peptides either at the N- and C-terminal ends (exopeptidases) or within the polypeptide chain (endopeptidases). These enzymes exist in all living organisms and it is estimated there are more than 66,000 different proteases [1, 2]. They are either localized within a cell, on or in the cell membrane, or secreted from the cell into the extracellular space [3]. Proteases are important for an organisms’ survival [4, 5]. Dysregulation of one or several proteases in humans and other vertebrates is associated with disease, and proteases are often involved when microorganisms invade a host [6–9]. Hence, proteases are important targets for therapeutic intervention [10, 11]. Proteases are classified into eight different classes/clans based on the amino acid or prosthetic group involved in the catalytic reaction. These classes are aspartic (A), cysteine (C), glutamic (G), metallo (M), aspargine (N), mixed (P), serine (S), and threonine (T) proteases (Merops database) [12]. There are more than 566 human and 644 murine proteases, of which 273/341 are secreted, 277/283 are intracellular, and 16/16

Davide Vigetti and Achilleas D. Theocharis (eds.), The Extracellular Matrix: Methods and Protocols, Methods in Molecular Biology, vol. 1952, https://doi.org/10.1007/978-1-4939-9133-4_16, © Springer Science+Business Media, LLC, part of Springer Nature 2019

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are intramembraneous, respectively. The majority of human and murine proteases are of the metallo, serine, and cysteine types, where most of the metallo- and serine proteases are secreted while the cysteine proteases are mainly localized within the cell [3]. Proteases induce an irreversible change of the substrate they process. Hence, most proteases are tightly regulated at the transcriptional, posttranscriptional, translational, and/or the posttranslational level. These enzymes are synthesized in an inactive pro-form, and are activated either within the cell or in the extracellular space. Once activated, their activity is regulated by protease inhibitors that bind either reversibly or irreversibly to the enzyme [6, 9, 13, 14]. To explore the physiological or pathological roles of proteases, it is important to be able to detect and precisely localize them in a tissue, to differentiate between inactive and active forms, as well as to quantify and determine the nature of the enzyme that degrades a given substrate. Active proteases can be determined in tissue by in situ substrate zymography. Here we present a protocol for real-time gelatin zymography, which is very useful for the detection of active gelatinases in tissue extracts. There are several proteases that degrade gelatin (denatured collagen), which include the two matrix metalloproteases MMP-2 and MMP-9 [15]. Other enzymes are serine proteases like trypsin, plasmin, and matriptase, as well as cysteine proteases such as cathepsin L [16–19]. In this protocol we focus on proteases that function around a neutral pH, but the protocol can be slightly changed for the detection of proteases with an optimal activity at slightly acidic pH (see Note 1). Proteases can be extracted from both unfixed and fixed tissue, but independent of the extraction method much larger amount of proteins is extracted from unfixed tissue compared to fixed tissue [20]. We have previously extracted proteins from kidney, liver, tongue, and heart tissue by mincing it in a tissue homogenizer with steel bullets in various extraction solutions including: 1. 0.25% Triton X-100 (v/v), 10 mM CaCl2 in Milli-Q water. 2. 10% DMSO (v/v), 10 mM CaCl2 in Milli-Q water. 3. Zymography loading buffer (1
): 0.05 M Tris–HCl, pH 6.8, 10% glycerol, 2.0% SDS, 0.05% bromophenol blue. 4. 1.0 M NaCl, 10 mM CaCl2 in Milli-Q water. The extracted proteins can be analyzed by different methods such as Western blotting, SDS-PAGE along with mass spectrometry, or gelatin SDS-PAGE zymography. The latter method reveals both the activity and the molecular size of the gelatin-degrading proteases. For MMPs, this method detects both the inactive pro-form and the active form of the protease because pro-MMPs refold and auto-activate when SDS is washed away after the electrophoresis. During the following incubation period (usually

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37  C) proteases can degrade the gelatin incorporated in the SDS gel. Even though gelatin SDS-PAGE zymography gives information of the gelatin-degrading enzymes present in the tissue, most reversible inhibitors will be dissociated from the proteases during SDS-PAGE, and hence it is not possible to conclude that the proteases detected are active in situ. Therefore, gelatin SDS-PAGE zymography and gelatin in situ zymography are complementary techniques for the characterization of gelatindegrading enzymes in a tissue. For both in situ and SDS-PAGE gelatin zymography, various types of inhibitors can be used to determine the type of protease responsible for the substrate degradation (see Note 2). When extracting gelatinolytic enzymes from a tissue, other proteins will follow. These non-gelatinolytic proteins can overlap and mask the gelatinolytic bands when running an ordinary gelatin SDS-PAGE gel. To avoid this problem, real-time SDS-PAGE zymography can be run instead. In contrast to normal gelatin zymography, real-time gelatin zymography is based on the use of fluorescence-labeled gelatin. Thus, it is not necessary to stain the gel after electrophoresis and hence other proteins in an extract will not interfere with the detection of gelatinolytic proteases. Another advantage is that it is possible to follow the development of gelatin degradation in real time by observing the gel under UV light (see Note 3). In this protocol, we will describe an easy, fast, and cheap method for labeling gelatin with the dye, 2-methoxy-2,4-diphenyl-3(2H)-furanone (MDPF). The dye reacts rapidly with primary amines and forms highly fluorescent and stable fluorophores (λex ¼ 385 nm and λem ¼ 480 nm), while the unbound dye and its hydrolysis products are nonfluorescent. The concentration of bound MDPF to gelatin can be determined by measuring the absorbance of the labeled gelatin at 385 nm, using the extinction coefficient, ε385nm  6500/M/cm. This protocol is based on MDPF labeling of triple-helical collagen [21]. Figure 1 shows a comparison between real-time gelatin zymography using MDPF-labeled gelatin and traditional normal gelatin zymography. Previously, Hattori et al. have described real-time zymography and real-time reverse zymography using FITC-labeled collagen, casein, and BSA [22]. The real-time zymography method presented here using MDPF-labeled gelatin can easily be converted to real-time reverse gelatin zymography by simply incorporating an appropriate amount of an enzyme like MMP-9, MMP-2, or trypsin in the SDS-PAGE gel along with the MDPF-gelatin (see Note 4). We have used this real-time reverse gelatin zymography method in several of our studies to detect the expression of tissue inhibitors of MMPs (TIMPs) [23–25].

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Fig. 1 Comparison of normal and real-time gelatin zymography. Kidney (K) and liver (L) samples from BALB/c mice were extracted either with zymography loading buffer (1
) (ZLB) or with 10% DMSO and 10 mM CaCl2 in Milli-Q water (DMSO). All samples were applied to the gels undiluted. (a) Normal gelatin zymography. Gel was incubated for approximately 17 h at 37  C in developing buffer and thereafter stained with Coomassie Brilliant Blue R-250. (b–e) Real-time gelatin zymography of the same samples as in (a) using MDPF-labeled gelatin. Pictures were taken after the gel had been washed and incubated in developing buffer for 2.5 h (b), 5 h (c, d), and 8.5 h (e) at 37  C. The pictures were obtained using a transilluminator (302 nm) and an ethidium bromide emission filter (transmit light between 580 and 630 nm) in front of the camera lens. The exception is for (c), where a transillumination converter plate (convert 302 nm to 365 nm) was used along with an emission filter in front of the camera lens that emits light between 465 and 495 nm. Standard (St) is a mixture of pro-MMP-9 (92 kDa), active MMP-9 (83 kDa) from THP-1 cells, and pro-MMP-2 (72 kDa) and active MMP-2 (62 kDa) from human skin fibroblasts. In normal zymography (a), gelatinase activity is shown as transparent zones in the dark-stained gelatin gel. In real-time zymography (b–e), gelatinase activity is shown as dark bands in the light fluorescence-emitting MDPF-gelatin gel

2

Materials

2.1 Reagents and Buffers

1. Gelatin (type A, porcine skin). 2. 2-Methoxy-2,4-diphenyl-3(2H)-furanone (MDPF). 3. Acetone. 4. Acrylamide/bis-acrylamide (29:1) (40% solution) (see Note 5). 5. Sodium dodecyl sulfate (SDS) (see Note 6). 6. Bromophenol blue. 7. N,N,N0 ,N0 -Tetramethylethylenediamine (TEMED). 8. 10% Ammonium persulfate (APS) in Milli-Q water (stored at 4  C). 9. Optional: Enzyme inhibitors such as ethylenediaminetetraacetic acid (EDTA), galardin, pefabloc. 10. 50 mM Sodium tetraborate, pH 9.0. 11. Separating gel buffer: 1.5 M Tris–HCl, pH 8.8, 0.4% SDS (see Note 6). 12. Stacking gel buffer: 0.5 M Tris–HCl, pH 6.8, 0.4% SDS.

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13. Electrophoresis buffer: 25 mM Tris–HCl, 190 mM glycine, pH 8.3, 0.1% SDS. 14. Zymography loading buffer (5
): 0.25 M Tris–HCl, pH 6.8, 50% glycerol, 10% SDS, 0.5% bromophenol blue (see Note 7). 15. Wash buffer: 2.5% Triton X-100 in Milli-Q water. 16. Developing buffer: 0.05 M Tris–HCl, pH 7.5, 0.2 M NaCl, 5 mM CaCl2, 0.02% Brij-35. 2.2

Equipment

1. Imaging system with a transilluminator (with 302 nm UV or 365 nm long-wave UV) and a camera with an emission filter in front of the lens. 2. Heating cabinet (37  C). 3. Vertical electrophoresis equipment with cooling. 4. Power supply. 5. Gel-casting system for polyacrylamide gels, including glass and alumina plates with appropriate spacer and well combs. 6. Optional: Handheld UV lamp (we use Model UVL-21, longwave UV 366 nm; UVP Inc.). 7. Optional: Converting plate with excitation filter converting UV 302 nm to UV 365 nm (Ultra-Violet Products LTD). 8. Optional: Emission filter that transmits light in the region 465–495 nm (Ultra-Violet Products LTD).

3

Methods

3.1 Labeling of Gelatin with MDPF

All solutions and steps containing MDPF should be protected from light. 1. Dissolve 37.5 mg gelatin in 5 mL 50 mM sodium tetraborate buffer pH 9.0. Warm the solution until the gelatin is dissolved, and then allow the solution to cool down to room temperature. Use a magnet for stirring. (To label larger amounts of gelatin, see Note 8.) 2. Dissolve 2 mg of 2-methoxy-2,4-diphenyl-3(2H)-furanone (MDPF) in 3 mL of acetone. 3. Add the MDPF solution slowly to the gelatin solution (room temperature; should take about 1 h), with continuous stirring. When all MDPF is added, keep on stirring for 1 h at room temperature. 4. Dialyze the labeled gelatin against 2
1 L of Milli-Q water at 4  C. This is to remove unbound MDPF and buffer. 5. Lyophilize the dialyzed MDPF-labeled gelatin (see Note 9).

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6. Dissolve the lyophilized MDPF-labeled gelatin in 1.875 mL Milli-Q water. This gives a gelatin concentration of approximately 2%. 7. Store in the dark in small batches. For long-time storage, use a freezer. 3.2 Preparation of MDPF-Gelatin SDSPAGE Separating Gel

1. The amounts given are for a separating gel with the following dimensions: (0.75 mm
8.2 cm
5.5 cm), i.e., a total volume of approximately 4.5 mL for 7.5%, 10%, and 13% polyacrylamide gels. A 7.5% gel is ideal for enzymes with a molecular size of 90 kDa or larger, while a 10% gel is best suited for enzymes with molecular sizes between 40 and 90 kDa and a 13% gel for enzymes with molecular sizes below 40 kDa (see Note 9). Solutions

PAGE 7.5% PAGE 10% PAGE 13%

1. Separating gel buffer (mL)

1.12

1.12

1.12

2. 2% MDPF-gelatin (μL)

225

225

225

3. Milli-Q water (mL)

2.266

1.974

1.625

4. 40% Acrylamide (mL)

0.874

1.166

1.515

5. TEMED (μL)

7.0

7.0

7.0

6. 10% APS (μL)

15

15

15

Mix the compounds in a plastic tube. Once the TEMED and APS have been added, the polymerization starts. Mix carefully and load the solution between the glass and alumina plate in a hand-cast polyacrylamide gel-casting system. Put gently a thin layer of Milli-Q water at the top of the loaded mixture and allow the acrylamide to polymerize (15–20 min). Protect the polymerizing gel from light! 3.3 Preparation of SDS-PAGE Stacking Gel

1. The amount is for a stacking gel with the following dimensions (0.75 mm
8.2 cm
2.5 cm), i.e., a total volume of approximately 1.5 mL. We give the contents of gels with 4% and 5% polyacrylamide. Solutions

PAGE 4%

PAGE 5%

1. Stacking gel buffer (mL)

0.186

0.186

2. Milli-Q water (mL)

1.145

1.106

3. 40% Acrylamide (μL)

155

194

4. TEMED (μL)

4.0

4.0

5. 10% APS (μL)

8.0

8.0

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Remove the water and un-polymerized acrylamide over the separating gel. Mix the compounds in a plastic tube. Once the TEMED and APS have been added, the polymerization starts. Mix carefully and load the solution on top of the SDS-PAGE separating gel. Put a well-forming comb on top, between the glass and alumina plate, and allow the acrylamide to polymerize (15–20 min). If you also expect to detect gelatinases with molecular sizes larger than 300 kDa, include also MDPF-gelatin in the stacking gel (see Note 10). Protect from light. 3.4 Electrophoresis and Visualization

1. Mount the gel plates on the electrophoresis apparatus. 2. Fill the lower and upper electrophoresis tray with electrophoresis buffer. 3. Mix 10 μL of samples and controls with 2.5 μL of loading buffer (5
). Notice, no boiling! 4. Apply 8 μL of the non-heated samples and controls onto the gel. 5. Start the cooling of the electrophoresis apparatus. 6. Make sure that the gel is protected from light! 7. Start the electrophoresis (constant current 20 mA/gel). 8. Stop the electrophoresis when the tracking dye has entered the bottom of the gel. 9. Remove the gel from the apparatus. 10. Remove the gel from the casting plates. Use a spatula. 11. Wash the gel 2
30 min in 50 mL wash buffer (the gel must be protected from light). 12. Add 50 mL of developing buffer (incubate at 37  C in the dark). 13. The degradation of gelatin can be visualized and photographed under UV light (366 or 302 nm) at appropriate time points. Gelatinases are detected as dark bands against the fluorescent MDPF-gelatin background.

4

Notes 1. To detect gelatin-degrading enzymes with a low pH optimum, use an appropriate developing buffer. O’Grady et al. [21] showed that the overall spectral pattern and the excitation and emission maxima were the same at pH 4.0 and 7.5. 2. Various types of inhibitors can be included in the wash and developing buffers to determine the type of protease responsible for the substrate degradation. For instance, 10 mM EDTA will inhibit metalloproteases, 1 μM galardin (Gm6001) matrix

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metalloproteases, and 1 mM pefabloc serine proteases. In the case of serine proteases, the sample can be incubated for 15 min with 1 mM pefabloc before the sample is mixed with 5
loading buffer and applied to the gel well, as this is an irreversible inhibitor. 3. Ideal is to use a UV lamp with long-range UV (366 nm). However, it is also possible to use a UV lamp with an excitation wavelength of 302 nm (see Fig. 1 in the introduction). 4. To perform real-time reverse gelatin zymography, add an appropriate amount of protease (recombinant, nonrecombinant, or cell-conditioned medium containing only the gelatin-degrading protease of interest) to the separating gel (see Subheading 3.2), and reduce the amount of Milli-Q water accordingly. We have used conditioned medium from PMA-stimulated THP-1 cells which contained proMMP-9 (92 kDa) and TIMP-1 (28 kDa) [23–25]. To allow detection of TIMPs (Mr 20–28 kDa) in samples, it was necessary to perform pre-electrophoresis of the gel before the samples were loaded, in order to remove the TIMP-1 incorporated into the gel. 5. Due to health security reasons, we use a commercial 40% acrylamide/bis-acrylamide solution in the preparation of all polyacrylamide gels. 6. Due to health security reasons, we use a commercial 20% SDS solution in the preparation of all buffers that contain SDS. 7. Due to health security reasons, when we make the zymography loading buffer (5
) we do not weigh the SDS powder, but use a commercial solution of 20% SDS and hence it is not possible to use 50% glycerol. Instead, we use 20% (w/v) sucrose. 8. For MDPF labeling of larger amounts of gelatin, adjust the amount of tetraborate buffer to obtain the same concentration of gelatin as step 1 in Subheading 3.1. Increase the amount of MDPF and acetone correspondingly. Also, adjust the volume of Milli-Q water during dialysis. 9. To use dialyzed MDPF-gelatin that has not been lyophilized, measure the volume of the dialyzed sample and calculate the concentration of MDPF-gelatin. Based on this, use an appropriate volume to obtain a concentration of 0.1% of MDPFgelatin in the separating gel. 10. Add 150 μL of 2% MDPF-gelatin and reduce the amount of Milli-Q water accordingly to get 0.2% of MDPF gelatin in the stacking gel. To avoid a dark zone between the separation and stacking gel due to a reduced amount of MDPF-gelatin, be careful to remove all water and un-polymerized acrylamide over the separating gel before the stacking gel is applied.

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Acknowledgments We are grateful to Dr. Kristin Andreassen Fenton (Faculty of Health Sciences, Department of Medical Biology, UiT-The Arctic University of Norway, Tromsø, Norway) for the gift of kidney and liver homogenates from a BALB/c mouse and senior engineer Eli Berg (Faculty of Health Sciences, Department of Medical Biology, UiT-The Arctic University of Norway, Tromsø, Norway) for performing the zymography experiments. We also would thank Rod Wolstenholme (Faculty of Health Sciences, UiT-The Arctic University of Norway, Tromsø, Norway) for help with the figure. References 1. Artenstein AW, Opal SM (2011) Proprotein convertases in health and disease. N Engl J Med 365:2507–2518 2. Winberg JO (2012) Matrix Proteinases: biological significance in health and disease. In: Karamanos NK (ed) Extracellular matrix: pathobiology and signaling. De Gruyter, Berlin, pp 230–238 3. Overall CM, Blobel CP (2007) In search of partners: linking extracellular proteases to substrates. Nat Rev Mol Cell Biol 8:245–257 4. Quiros PM, Langer T, Lopez-Otin C (2015) New roles for mitochondrial proteases in health, ageing and disease. Nat Rev Mol Cell Biol 16:345–359 5. Ricard-Blum S, Vallet SD (2016) Proteases decode the extracellular matrix cryptome. Biochimie 122:300–313 6. Hadler-Olsen E, Fadnes B, Sylte I et al (2011) Regulation of matrix metalloproteinase activity in health and disease. FEBS J 278:28–45 7. Maeda H (1996) Role of microbial proteases in pathogenesis. Microbiol Immunol 40:685–699 8. Tokito A, Jougasaki M (2016) Matrix metalloproteinases in non-neoplastic disorders. Int J Mol Sci 17. https://doi.org/10.3390/ ijms17071178 9. Turk V, Stoka V, Vasiljeva O et al (2012) Cysteine cathepsins: from structure, function and regulation to new frontiers. Biochim Biophys Acta 1824:68–88 10. Brotz-Oesterhelt H, Sass P (2014) Bacterial caseinolytic proteases as novel targets for antibacterial treatment. Int J Med Microbiol 304:23–30 11. Gialeli C, Theocharis AD, Karamanos NK (2011) Roles of matrix metalloproteinases in cancer progression and their pharmacological targeting. FEBS J 278:16–27

12. Rawlings ND, Barrett AJ, Thomas PD et al (2018) The MEROPS database of proteolytic enzymes, their substrates and inhibitors in 2017 and a comparison with peptidases in the PANTHER database. Nucleic Acids Res 46: D624–D632 13. Gaffney J, Solomonov I, Zehorai E et al (2015) Multilevel regulation of matrix metalloproteinases in tissue homeostasis indicates their molecular specificity in vivo. Matrix Biol 44–46:191–199 14. Overall CM, Lopez-Otin C (2002) Strategies for MMP inhibition in cancer: innovations for the post-trial era. Nat Rev Cancer 2:657–672 15. Malla N, Sjoli S, Winberg JO et al (2008) Biological and pathobiological functions of gelatinase dimers and complexes. Connect Tissue Res 49:180–184 16. Heussen C, Dowdle EB (1980) Electrophoretic analysis of plasminogen activators in polyacrylamide gels containing sodium dodecyl sulfate and copolymerized substrates. Anal Biochem 102:196–202 17. Lin CY, Anders J, Johnson M et al (1999) Molecular cloning of cDNA for matriptase, a matrix-degrading serine protease with trypsinlike activity. J Biol Chem 274:18231–18236 18. Spens E, Haggstrom L (2005) Protease activity in protein-free NS0 myeloma cell cultures. In Vitro Cell Dev Biol Anim 41:330–336 19. Winberg JO, Gedde-Dahl T (1986) Gelatinase expression in generalized epidermolysis bullosa simplex fibroblasts. J Invest Dermatol 87:326–329 20. Hadler-Olsen E, Kanapathippillai P, Berg E et al (2010) Gelatin in situ zymography on fixed, paraffin-embedded tissue: zinc and ethanol fixation preserve enzyme activity. J Histochem Cytochem 58:29–39

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21. O’grady RL, Nethery A, Hunter N (1984) A fluorescent screening assay for collagenase using collagen labeled with 2-methoxy-2,4diphenyl-3(2H)-furanone. Anal Biochem 140:490–494 22. Hattori S, Fujisaki H, Kiriyama T et al (2002) Real-time zymography and reverse zymography: a method for detecting activities of matrix metalloproteinases and their inhibitors using FITC-labeled collagen and casein as substrates. Anal Biochem 301:27–34 23. Loennechen T, Mathisen B, Hansen J et al (2003) Colchicine induces membrane-

associated activation of matrix metalloproteinase-2 in osteosarcoma cells in an S100A4-independent manner. Biochem Pharmacol 66:2341–2353 24. Malla N, Berg E, Theocharis AD et al (2013) In vitro reconstitution of complexes between pro-matrix metalloproteinase-9 and the proteoglycans serglycin and versican. FEBS J 280:2870–2887 25. Mathisen B, Lindstad RI, Hansen J et al (2003) S100A4 regulates membrane induced activation of matrix metalloproteinase-2 in osteosarcoma cells. Clin Exp Metastasis 20:701–711

Chapter 17 In Vitro Spheroid Sprouting Assay of Angiogenesis Fatema Tuz Zahra, Efrossini Choleva, Md Sanaullah Sajib, Evangelia Papadimitriou, and Constantinos M. Mikelis Abstract Angiogenesis is a well-coordinated physiological process that leads to new blood vessel formation. Physiologically, angiogenesis is more prominent during development and wound healing and its dysregulation drives or is related to several diseases, including cancer. The endothelial cells are the main regulators of the angiogenic process, and thus the angiogenic outcome is assessed based on the effect on endothelial cell functions. Several in vitro and in vivo techniques have been developed to assess the effect of various factors on angiogenesis. Compared to the in vivo techniques, the in vitro techniques are considered less physiologically relevant. This has been partially overcome by the development of 3-dimensional (3D) in vitro models, one of which is the spheroid assay or 3D sprouting assay that exploits the effect of the extracellular matrix to endothelial cell functions. This chapter focuses on the description of the spheroid assay and mentions the variations and potential applications this assay can have. Key words Angiogenesis, Endothelial cells, Collagen, Spheroids, Sprouts

1

Introduction In vascular research, the in vivo models are considered more physiologically relevant than the in vitro ones, similar to other science fields. The main reasons are the involvement of a live organism, defined by the participation of different cell types, which can affect the biological outcome, as well as the presence of the 3-dimensional microenvironment, which regulates and sometimes defines the final outcome. Although the in vivo models bear high physiological relevance, their main disadvantages remain the reduced reproducibility, due to variation within species, and the high cost. Therefore, the in vitro models are routinely used for assessment of angiogenesis. The main advantages of the in vitro models remain the ability to analyze the effect on a particular cell type, allowing molecular pathway analysis in the specific cell type, and the increased reproducibility, which also provides the potential to scale up the assay for screening purposes [1, 2]. Compared to the 2D traditional

Davide Vigetti and Achilleas D. Theocharis (eds.), The Extracellular Matrix: Methods and Protocols, Methods in Molecular Biology, vol. 1952, https://doi.org/10.1007/978-1-4939-9133-4_17, © Springer Science+Business Media, LLC, part of Springer Nature 2019

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angiogenesis models, the 3D in vitro models are considered more proximal to the human physiology, without compromising the advantages the in vitro models have. The 3D in vitro model for angiogenesis is the spheroid or 3D sprouting assay, which was developed around a decade ago [1]. This assay can be performed with endothelial cells of any origin, such as human umbilical vein endothelial cells (HUVECs), human dermal microvascular endothelial cells (HDMECs), or human dermal lymphatic endothelial cells (HDLECs) [1]. Furthermore, all basic endothelial functions (proliferation, migration, sprout formation) are required for the sprouting process; therefore, it can be successfully applied for screening purposes on angiogenesis evaluation [2]. Although the description within this chapter is focused on the spheroids as an in vitro technique, it has been developed to be used as an in vivo model as well [1].

2 2.1

Materials Reagents

1. Human umbilical vein endothelial cells (HUVECs) (see Note 1). 2. Medium 199 (M199), stored at 4
C. 3. Fetal bovine serum (FBS), aliquoted and stored at 20
C (see Note 2). 4. Endothelial cell growth supplement (ECGS), stored as lyophilized powder at 4
C or at 20
C for long-term storage. 5. Heparin (5000 U/mL stock), stored at room temperature (RT). 6. Penicillin/streptomycin (P/S antibiotic cocktail), stored at 4
C. 7. Trypsin/EDTA solution, stored at 4
C. 8. Methyl cellulose. 9. Collagen type I, rat-tail high concentrate (8–11 mg/mL) in 0.02 N acetic acid, stored at 4
C (see Note 3). 10. 1 M HEPES (stored at 4
C). 11. 0.2 M NaOH (stored at RT). 12. Phosphate-buffered saline (PBS) (137 mM NaCl, 2.7 mM KCl, 10 mM Na2HPO4, 1.8 mM KH2PO4, pH 7.4), sterile, stored at RT. 13. Paraformaldehyde (PFA) solution 4% in PBS, stored at 4
C. 14. 0.1% Acetic acid in water. 15. 0.2% Triton X-100 in PBS. 16. 3% Bovine serum albumin (BSA) in PBS.

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17. Phalloidin (for actin filament staining): 2 μL in 500 μL of 3% BSA in PBS. 18. Hoechst for nuclei staining: Dilute 1:2000 in PBS before use. 2.2 HUVEC Growth Medium Preparation

Resuspend ECGS into Medium 199 (M199) and incubate for 30 min at 37
C. Pass the medium through a 0.22 μm-pore filter inside the cell culture flow hood. Add heparin to 5 U/mL final concentration, antibiotic cocktail to a 1 final concentration, and FBS to 15% final concentration. Preserve under sterile conditions at 4
C.

2.3 Methocel Preparation

Autoclave 3 g methyl cellulose with magnetic stirrer in a 500 mL glass beaker covered with aluminum foil. Add 125 mL of HUVEC starvation media (M199) inside the cell culture flow hood and stir for 20 min covered. Add 125 mL more of M199 in the cell culture flow hood and stir overnight at 4
C. Aliquot the solution evenly to 50 mL Falcon tubes and centrifuge at 3500  g for 3 h at 4
C. The final concentration should be 1.2% Methocel in starvation media (M199). The solutions can be kept at 4
C for ~3 months (see Note 4).

2.4 Preparation of 10% FBS in PBS

Calculate volumes according to the numbers of plates with spheroids used (10 mL per plate) and 10 mL more for a final wash.

2.5 Methocel/FBS/ Antibiotics (Pen/Strep) Preparation

Methocel/FBS/antibiotics (Pen/Strep): 80% Methocel + 20% FBS (0.5 mL per well of 24-well plate). Add in a tube the calculated amount of FBS and antibiotics (P/S, 1–100 of FBS), and then add the calculated amount of Methocel. Mix it once and leave it for the bubbles to come to the surface.

2.6 Preparation of 20% Methocel in HUVEC Growth Medium

Add 20% v/v Methocel with 80% v/v HUVEC growth medium (i.e., 2 mL Methocel and 8 mL HUVEC growth medium for 10 mL final volume) in a conical sterile polypropylene centrifuge tube. Mix gently with serological pipettes. Calculate the required volume of 20% Methocel based on the number of cells (400 cells correspond to 25 μL of the solution: i.e., 10 mL for 160,000 cells). Always prepare for 2–5 gels more than the experimental number.

2.7

Equipment

1. Plastic square petri dishes, sterile. 2. 15 mL Conical sterile polypropylene centrifuge tubes. 3. 50 mL Conical sterile polypropylene centrifuge tubes. 4. Centrifuge for eukaryotic cell culture. 5. Laminar flow hood. 6. CO2 gas incubator. 7. Pipettes for volumes 10–1000 μL.

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8. Pipette tips for volumes 10–1000 μL. 9. Multi-micropipette, 12-channel, 10–100 μL. 10. Cell culture petri dishes, sterile. 11. Cell counting chamber Neubauer haemocytometer. 12. Sterile disposable solution basin. 13. 24-Well tissue culture plates. 14. Inverted microscope. 15. Fluorescent microscope.

3

Methods

3.1 Spheroid Preparation and Embedding

1. Calculate the required number of cells, based on the number of experimental groups used in the specific experiment (see Note 5).

3.1.1 Day 1: Spheroid Preparation

2. Aspirate HUVEC growth medium of a 100 mm HUVEC culture plate of 70–80% confluency. 3. Wash with sterile PBS and aspirate. 4. Repeat once. 5. Add 1 mL trypsin/EDTA solution and incubate at 37
C for 1 min. 6. Stop the reaction by adding 3 mL PBS containing 10% FBS (v/v) on the cells. 7. Pipette the cells into a 15 mL centrifuge tube and centrifuge them for 5 min 150  g, RT. 8. Discard the supernatant and resuspend the cells in 2–3 mL of HUVEC growth medium. 9. Count the cells. This step can be performed through an automated cell counter or manually through a cell counting chamber. 10. Take the calculated number of cells in a 50 mL centrifuge tube and spin them, followed by resuspending them to 20% Methocel in HUVEC growth medium. Mix them gently using a 25 mL pipette. 11. Distribute cell suspension in 25 μL drops (see Note 6) on the lid of square petri dishes using a sterile disposable solution basin and a multichannel pipette (12-channel pipette). Close the petri dish with lid (so that the cell droplets will hang) and incubate at 37
C for 24 h.

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3.1.2 Day 2: Spheroid Harvesting, Resuspension, and Embedding into Collagen Gels

1. Check under the phase-contrast microscope whether the cells have formed evenly rounded spheroids. Proceed only if the shape and size of the spheroids are round and uniform, respectively.

Spheroid Harvesting

2. In a 24-well plate, fill the extra wells with 1 mL PBS and incubate at 37
C for at least 20 min. This is required, in order to increase the temperature of the plate, so that the gel will polymerize immediately, preventing the spheroids to concentrate in the bottom of the wells upon embedding. 3. Harvest spheroids with 10% FBS in PBS (10 mL/plate): Adjust the suction speed of the pipette gun to the lowest. Collect the spheroids with 5 mL 10% FBS in PBS with 25 mL pipette and transfer them into the 50 mL Falcon tubes. Smooth movements are required to preserve the integrity of the spheroids. Wash the plate once again with another 5 mL 10% FBS in PBS, as the spheroids tend to stick to the dish. If required, use 5 mL more to collect all spheroids. 4. Centrifuge at 100  g for 5 min at 25
C. 5. Aspirate most of the supernatant (should be noted that not all supernatant should be collected, because spheroids can be lost, as the pellet is not clearly visible). 6. Pour Methocel/FBS/antibiotics (calculate as 0.5 mL of Methocel/FBS/antibiotics per well) to the tube containing the spheroids with slow motion. Do not mix! The liquid must be dropped from the pipette very slowly.

Collagen Gel Preparation (For Quantity See Note 7)

1. In a 50 mL Falcon tube add acetic acid, collagen and M199 in a 4:4:1 ratio. The color of the solution will turn to yellow. 2. Add 0.2 M NaOH slowly and dropwise. After you add each drop shake it mildly and shortly (on ice) to see if the color changes. Do this till the color starts becoming salmon pink. At that point, add one or two drops of NaOH solution more. The color should turn back to pink. 3. Then, add 1 M HEPES drop by drop till the pink color turns salmon pink again. Leave the collagen mix on ice and go to the next step quickly. 4. Mix the collagen mix with Methocel/FBS/antibiotics/spheroids (resuspended spheroids) 1:1. This step needs to be done in a way to ensure minimizing bubble generation and relatively fast. 5. Add 1 mL from the mixture to each well and incubate for 30 min at 37
C for polymerization. 6. Stimulate the cells with 100 μL of Medium 199 with or without growth factors and incubate at 37
C for 24 h (see Note 8).

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3.1.3 Day 3: Sample Fixation

3.2 Phalloidin and Hoechst Staining of Spheroids

1. Add 1 mL of 4% PFA to each well. Do not wipe out the PFA. 2. Take images of the spheroids with inverted microscope and quantify number of sprouts and sprout length, using the ImageJ software. Upon PFA fixation, the spheroids can be either visualized directly (above and Fig. 1) or stained with phalloidin for optimal identification of the cell sprouts (Fig. 2). Staining can be performed from 4 h after fixation to a few days later. The samples should be kept at 4
C and liquid should be regularly added to avoid dryness.

Fig. 1 Bright-field image of an endothelial spheroid. This spheroid consists of 400 endothelial cells. The arrows highlight two newly formed sprouts

Fig. 2 Representative fluorescent image of a spheroid by confocal microscope. The blue color corresponds to endothelial cell nuclei and the sprouts can be easily identified by the actin staining (red color)

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The staining takes place through the following process:

1. Remove 4% PFA from the top of the gel. 2. Wash with PBS for 5 min at RT. 3. Repeat the wash once. 4. Permeate the cells with 0.2% Triton X-100 for 10 min (500 μL per well). 5. Wash with PBS for 5 min at RT. 6. Repeat the wash once. 7. Block with 3% BSA in PBS for 1 h at RT (500 μL per well). 8. Remove liquid from top of the gels carefully (do not touch the gel). 9. Add phalloidin (2 μL in 500 μL 3% BSA in PBS per well) and incubate overnight at 4
C. 10. Wash with 500 μL to 1 mL of PBS for 10 min at RT. 11. Repeat the wash once. 12. Add Hoechst (500 μL) and incubate for 10 min at RT. 13. Wash with 500 μL to 1 mL of PBS for 10 min at RT. 14. Repeat the wash once. 15. Store at 4
C, covered with aluminum foil. 16. Obtain images with fluorescent microscope.

4

Notes 1. This protocol has been performed with human umbilical vein endothelial cells (HUVECs). HUVECs are used at passages 1–6. For any other primary endothelial cell types, the appropriate media have to be used and the conditions may be optimized. 2. FBS should be heat-inactivated at 56
C for 30 min, upon thawing. After heat inactivation, it should be kept at RT for around 15 min and then divided into aliquots under sterile conditions (cell culture hood) and stored at 20
C. 3. The collagen stock, if not purchased, should be in 1:2 or 1:1 dilution with 0.1% acetic acid, so that the final concentration reaches around 5 mg/mL. 4. As the sediment contains residual cellulose fibers, only the upper 45 mL from the 50 mL aliquots should be used for the experiment. 5. For the calculation of cells, the following rule can be helpful: 1 spheroid ¼ 400 cells ¼ 25 μL of medium. 1 gel ¼ 100 spheroids ¼ (100  400 cells) ¼ 40,000 cells/gel

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6. For the calculation of the volume of the 20% Methocel/80% HUVEC growth medium, where the cells will be resuspended to form spheroids, the following rule can be helpful: 1 spheroid ¼ 400 cells ¼ 25 μL of medium. 1 gel ¼ 100 spheroids ¼ (100  25 μL) ¼ 2.5 mL of medium/ gel 7. For collagen mix preparation, keep all reagents on ice. Dilute collagen 1:2 in 0.1% acetic acid. Calculate the amounts of reagents to add 0.5 mL of gel per well. For example, 5 mL of gel final volume will require 2 mL collagen stock (concentration 8–11 mg/mL), 2 mL 0.1% acetic acid, 500 μL M199, approximately 300 μL NaOH, and approximately 160 μL HEPES. 8. Important: The growth factor concentration should be calculated in a volume of 1100 μL (11 higher concentration than final in the 100 μL to be added; examples of growth factors used in spheroids are vascular endothelial growth factor (VEGF), basic fibroblast growth factor (bFGF), and angiopoietin-2 (Ang2)).

Acknowledgments Effrosini Choleva is recipient of a PhD scholarship from the State Scholarship Foundation in Greece (IKY) (Operational Program “Human Resources Development—Education and Lifelong Learning,” Partnership Agreement PA 2014-2020). Human umbilical vein endothelial cells (HUVEC) were isolated from human umbilical cords in accordance with the relevant guidelines and regulations from the Committee for Ethics in Research of the University of Patras or the Texas Tech University Health Sciences Center Institutional Review Board (IRB) under an IRB-approved protocol (IRB#A15-3891). Informed consent was obtained from all subjects. References 1. Laib AM, Bartol A, Alajati A, Korff T, Weber H, Augustin HG (2009) Spheroid-based human endothelial cell microvessel formation in vivo. Nat Protoc 4:1202–1215. https://doi.org/10. 1038/nprot.2009.96

2. Heiss M, Hellstro¨m M, Kale´n M, May T, Weber H, Hecker M, Augustin HG, Korff T (2015) Endothelial cell spheroids as a versatile tool to study angiogenesis in vitro. FASEB J 29:3076–3084. https://doi.org/10.1096/fj. 14-267633

Chapter 18 Matrigel Plug Assay for In Vivo Evaluation of Angiogenesis Pinelopi Kastana, Fatema Tuz Zahra, Despoina Ntenekou, Stamatiki Katraki-Pavlou, Dimitris Beis, Michail S. Lionakis, Constantinos M. Mikelis, and Evangelia Papadimitriou Abstract Matrigel is extracted from the Engelbreth-Holm-Swarm (EHS) mouse sarcoma in C57BL/6 mice, a tumor rich in extracellular matrix (ECM) proteins. It consists mainly of laminin (approximately 60%), collagen IV (approximately 30%), and nidogen-1/entactin (approximately 8%), while it also contains heparan sulfate proteoglycans, such as perlecan, other ECM proteins, as well as growth factors bound to the ECM. Matrigel mimics the physiological cell matrix and is the most commonly used matrix substrate to study in vitro and in vivo angiogenesis. Here, we describe the in vivo Matrigel plug assay and how it can be used for both qualitative and quantitative assessment of angiogenesis. Key words Angiogenesis, Endothelial cells, Matrigel, Matrigel plug, Vessels

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Introduction The in vivo Matrigel plug assay is widely used to assess the in vivo angiogenic potential or anti-angiogenic activity of different compounds in animal models. It is derived from C57BL/6 mice and is mostly used in this strain to avoid immunological reactions [1]. The test substance(s) can be suspended in the gel, which is liquid at 4
C and forms a solid gel at 37
C. The liquid gel is then injected subcutaneously in mice, where it instantly forms a solid plug allowing the slow release of the substance. At the end of the experiment, the plugs are removed and analyzed for the formation of blood vessels [1–4]. More recently, a more sophisticated and accurate method has been introduced, the so-called directed in vivo angiogenesis assay (DIVAA) [5]. The main principle is the same as in the traditional plug assay, but instead of using Matrigel plugs, silicone cylinders are filled with Matrigel and subsequently implanted subcutaneously into mice. Endothelial cells migrate inside the cylinder and form vessels.

Davide Vigetti and Achilleas D. Theocharis (eds.), The Extracellular Matrix: Methods and Protocols, Methods in Molecular Biology, vol. 1952, https://doi.org/10.1007/978-1-4939-9133-4_18, © Springer Science+Business Media, LLC, part of Springer Nature 2019

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Materials 1. Basement membrane matrix (Matrigel) or growth factorreduced basement membrane matrix (see Note 1). 2. 1 mL Single-use syringes with 25G needle. 3. Eppendorf tubes. 4. Trimmer to shave mouse flanks when needed, e.g., when mice of the C57BL/6 background are used (see Note 2). 5. C57BL/6 mice. 6. Inhalant isoflurane. 7. Ketamine/xylazine cocktail (through intraperitoneal injection): It contains 87.5 mg/kg ketamine and 12.5 mg/kg xylazine (anesthesia lasts approximately 20 min). 8. DMEM full medium: DMEM containing 10% fetal bovine serum and antibiotics. 9. PBS: NaCl 8 g/L, KCl 0.2 g/L, Na2HPO4 1.44 g/L, KH2PO4 0.24 g/L in distilled water. Adjust pH to 7.4 with HCl. Prior to use, the buffer is autoclaved for 20 min at 121
C. 10. TBS 10: Tris base 24.2 g/L, NaCl 80 g/L in distilled water. Adjust pH to 7.6 with HCl. Prior to use, the buffer is diluted ten times with distilled water. 11. 1 TBS, 0.05% (v/v) Tween-20 (TBS-T). 12. Mayer’s hematoxylin solution. 13. Eosin Y solution. 14. DPX (phthalate free) mounting medium (Electron Microscopy Sciences) or similar. 15. 3% or 30% bovine serum albumin (BSA) in PBS. 16. 2% or 4% formaldehyde in PBS: Dilute from a formaldehyde solution >34.5% wt. 17. 0.2% Triton X-100 and 200 mM glycine in PBS. 18. Biopsy cassettes. 19. Mounting medium. 20. FACS buffer: 1% BSA, 0.01% NaN3 in PBS. 21. Collagenase type II solution: Prepare 0.1% collagenase (1 mg/ mL) in CellSperse solution. Calculate for 1 mL/sample. 22. Dispase (1.88 units/mg) solution: Prepare 4 mg/mL of dispase in collagenase type II solution to have a final concentration of 0.8 U/mL. 23. DNase solution: Prepare 0.1% DNase in CellSperse solution and dilute 1:100 in dispase solution.

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24. Digestion solution: 0.1% Collagenase type II, 0.8 U/mL Dispase, and 0.001% DNase in CellSperse solution. 25. Staining kit for live/dead cells that contains a dye to determine cell viability by validation through UV laser. 26. Donkey anti-rat Alexa Flour 488 (see Note 3). 27. CD16/CD32 monoclonal antibody (see Note 3). 28. Rat anti-mouse CD31 (see Note 3). 29. CD16/CD32 monoclonal antibody (see Note 3). 30. Alexa Fluor 488-conjugated LYVE-1 (see Note 3). 31. PE-Cy7-conjugated CD31 (see Note 3). 32. Allophycocyanin (APC)-conjugated TER-119 (see Note 3). 33. APC-Cy7-conjugated CD45 (see Note 3). 34. eFluor 450-conjugated CD102 (see Note 3). 35. PE-conjugated fluorescent counting beads. 36. Rhodamine Griffonia simplicifolia lectin I: Dilute 1:200 in TBS-T prior to use. 37. Hoechst for nuclei staining: Dilute 1:2000 in PBS before use. 38. Draq 5 for nuclei staining: Dilute 1:1000 in PBS before use. 39. Mowiol 4-88: Add 2.4 g Mowiol to 6 mL glycerol. While mixing, add 6 mL distilled water and leave for 2 h at room temperature (RT). Then, add 12 mL 0.2 M Tris–HCl (pH 8.5). Incubate at 50–60
C until Mowiol dissolves. Filter, aliquot, and keep at 20
C. 40. Directed In vivo Angiogenesis Assay (DIVAA) Kit (see Note 4). 41. 100% Ethanol and ethanol solutions diluted in water. 42. Xylene. 43. Glass slides. 44. Cryotome. 45. Microtome. 46. FACS equipment. 47. Confocal microscope. 48. Fluorescent microscope. 49. Fluorescence plate reader.

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Methods

3.1 Matrigel Injection and Plug Formation

1. Thaw Matrigel (from 80
C) on ice at 4
C overnight. 2. Put all Eppendorf tubes, syringes, and needles on ice. This will inhibit Matrigel polymerization.

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3. Add 500 μL of Matrigel in a 1.5 mL Eppendorf tube and add the substance(s) to be tested. The Eppendorf tube should be kept on ice (see Notes 5 and 6). 4. Anesthetize each mouse immediately before Matrigel injection. The most common ways are using inhalant isoflurane or subcutaneously injected ketamine/xylazine. When inhalant isoflurane is used, 5% can induce anesthesia, while 1–3% is used for preserving the anesthetic effect during the procedure (see Note 7). Place 1–2 animals in the anesthesia inhalation chamber and seal top, ensure flow of isoflurane 500 mL/min with the vaporizer in 5%, and check the animals. The animals are put in special facemasks that provide 100–200 mL/min isoflurane and vaporizer in 2% for maintaining anesthesia for approximately 20 min. (After extracting facemasks, the animals are awake within 5 min.) 5. Shave the flank area of mouse (see Note 8) and inject Matrigel subcutaneously into the flank (slow injection). Allow it to solidify. Normally, both flanks of the same mouse are used, but this depends on the experiment. 6. Matrigel incubation in the mice takes place for 7–15 days. 7. After euthanizing mice (see Note 9), take the plugs out and put them onto 12-well plate filled with DMEM full medium till all plugs are collected (see Note 10). 8. Remove connective and adipose tissues surrounding the plug. 9. Briefly wash the plugs by incubating them into PBS. 10. Take pictures. Keep the same background for all plugs. Intensity of red color of the plug denotes angiogenesis levels (Fig. 1).

Fig. 1 Representative image of an extracted Matrigel plug. Levels of transparency, as well as intensity of the red color, are used as parameters for qualitative assessment

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11. Upon extraction, the plugs can be destined for sectioning (cryo-sectioning or paraffin sectioning) or cell isolation. 3.2 Preparation of Matrigel Plug for Cryo-Sectioning

1. Take liquid nitrogen in a box and drop the plug after extraction and washing (see Subheading 3.1, steps 7–9) in the liquid nitrogen for 20–30 s until bubbles come up. 2. Remove the plug from liquid nitrogen and store at 80
C. 3. Before doing the cryo-section, keep the samples out from 80
C to 20
C for 2–3 h. 4. Make 40 μm sections using cryotome. Transfer the sections on glass slides and store at 20
C.

3.3 Preparation of Matrigel Plug for Paraffin Sectioning

1. After extraction and washing (see Subheading 3.1, steps 7–9), fix the plugs in 2% formaldehyde for 24 h, at room temperature. 2. Wash with PBS and store in 70% ethanol. The plugs can be stored in ethanol for approximately 3 months. Change the ethanol occasionally for long-term storage. 3. Embed plugs in paraffin block: Put each plug in a biopsy cassette and mark by pencil. Embed twice for 5 min in 70%, 90%, 95%, and 100% ethanol solutions, followed by washing with xylene. (This dehydration process can be done in an automated tissue processor.) Exchange xylene with paraffin. This is done in an oven set for 54–58
C. Embed in fresh new paraffin and orient tissue as desired. Then, the block is put on an iced surface and is left to harden. 4. Cut thin slices (5 μm) of the paraffin-embedded plugs with a microtome. 5. Transfer the slices on glass slides and dry at 40
C for at least 2 min. 6. You can store the slides at room temperature for long time or move forward for staining.

3.4 Hematoxylin and Eosin (H&E) Staining of Matrigel Plug Paraffin Sections

1. Incubate the section slides in oven at 65
C for 30 min for paraffin evaporation. 2. Rehydrate the sections by dipping into the following solutions serially for 10 min each: xylene > xylene > xylene > 100% ethanol > 95% ethanol > 90% ethanol > 70% ethanol (under chemical fume hood). 3. Rinse the slides in cold tap water, keeping them in the bucket (2–3 min). 4. Dip the slides into Mayer’s hematoxylin solution for 20 min (nuclear staining). 5. Rinse the slides with hot running water for 10–15 min.

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Fig. 2 Representative image of a 5 μm thick Matrigel plug paraffin section. Cells and nuclei are stained with H&E. Matrigel is also stained (pale background color). Vessel lumens are characterized by absence of staining and marked with asterisks

6. Dip the slides in Eosin Y for 1–2 min (cytoplasmic staining). 7. Dehydrate the slides again with the reverse sequence (70% ethanol > 90% ethanol > 95% ethanol > 100% ethanol > xylene > xylene > xylene), 10 min each. 8. Mount with DPX mounting medium. 9. Take images under microscope [2] (Fig. 2). 3.5 Griffonia simplicifolia Lectin Staining of Matrigel Plug Paraffin Sections

There are several, well-characterized methods to quantify angiogenesis in sections from paraffin-embed tissues. A common method is immunostaining of endothelial cells for CD31 and direct measurement of the stained vessels under a light or fluorescent microscope. An alternative is staining the vessels with the endothelial specific lectin Griffonia simplicifolia that is conjugated to a fluorescent dye. Stained vessels can then be measured under a fluorescent microscope. Griffonia simplicifolia binds to endothelial cells of different species and thus, there is no need for tedious protocol adjustments related to the use of antibodies from different sources for different tissues/species. 1. Incubate the section slides at 37
C for at least 20 min for paraffin evaporation. 2. Rehydrate the sections by dipping into the following solutions serially as follows: xylene (37
C, 10 min) > xylene (RT, 10 min) > xylene (RT, 10 min) > 100% ethanol (RT, 10 min) > 96% ethanol (RT, 10 min) > 80% ethanol (RT,

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5 min) > 50% ethanol (RT, 5 min) (under chemical fume hood). 3. Rinse the slides in cold tap water, keeping them in the bucket (2–3 min). 4. Apply Griffonia simplicifolia solution directly onto the section. 5. Incubate for 1 h in the dark at RT. 6. Wash 5 with TBS-T for 5 min each. 7. Apply Draq5 for 5 min at 37
C to stain cell nuclei. 8. Mount with Mowiol (see Note 11). 9. Keep the slides at 4
C in the dark until observation. 10. Take images with fluorescent microscope. The images can be evaluated either qualitatively, based on the density of the vascular plexus, or quantitatively, based on the number of nuclei (blue color), number of sprouts, or size of diameter of highlighted vessels per image. 3.6 Immunofluorescent (CD31) Staining of Matrigel Plug Cryo-Sections

Immunostaining of endothelial cells for CD31 is the most widely used method to study angiogenesis, despite the effort required to find the proper antibody. Some antibodies work better in cryosections and this approach is also of choice in cases of co-staining of CD31 with other markers. 1. Wash the tissue sections on glass slide once with PBS (add dropwise carefully on slide). 2. Incubate the slide with 4% PFA in PBS for 20 min at RT (add dropwise carefully on slide). 3. Quick wash with PBS. 4. Repeat the quick wash with PBS. 5. Incubate in a 0.2% Triton X-100 and 200 mM glycine in PBS solution at RT for 15 min (tissue permeation). 6. Quick wash with PBS. 7. Repeat the quick wash with PBS. 8. Block the tissue with 3% BSA in PBS at RT for 2 h. 9. Incubate the slide with the rat anti-mouse CD31 primary antibody, 1:10 dilution in 3% BSA in PBS at 4
C overnight. 10. Quick wash with PBS. 11. Repeat the quick wash with PBS. 12. Wash with PBS for 5 min. 13. Repeat the wash with PBS for 5 min. 14. Add the donkey anti-rat Alexa Flour 488 secondary antibody, 1:500 in 3% BSA in PBS, and incubate in the dark at RT for 2 h. 15. Quick wash with PBS.

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Fig. 3 Representative image of a 40 μm thick Matrigel plug cryo-section. Endothelial cells are stained for CD31 (green) and nuclei are stained with Hoechst (blue). Vessel sprout is distinguished in the center and sections of bigger vessels are in the background

16. Repeat the quick wash with PBS. 17. Apply Hoechst for 10 min in the dark at RT. 18. Quick wash with PBS. 19. Repeat the quick wash with PBS. 20. Wash with PBS twice for 5 min. 21. Repeat the wash with PBS for 5 min. 22. One quick wash with H2O followed by mounting medium. 23. Keep in the dark for approximately 1 h for solidification, and store at 4
C. 24. Take images with fluorescent microscope (Fig. 3). The images can be evaluated either qualitatively, based on the density of the vascular plexus, or quantitatively, based on the number of nuclei (blue color), number of sprouts, or size of diameter of highlighted vessels per image. 3.7 Matrigel Plug Cell Isolation and Characterization by Flow Cytometry (FACS)

1. After euthanizing mice, take the plugs out and put them onto 12-well plate filled with DMEM full media till all plugs are collected (see Subheading 3.1, steps 1–7). 2. Remove connective and adipose tissues surrounding the plug. 3. Briefly wash the plugs by incubating to PBS. 4. Mince the plugs in smaller pieces by a blade.

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5. Prepare the digestion solution beforehand (see Note 12). 6. Put 1 mL per sample of the digestion solution in a 2 mL Eppendorf tube. 7. Incubate at 37
C with 500 rpm shaking to dissolve the Matrigel plug (it will take approximately 2 h, depending on the angiogenic growth in the plug). 8. When the plugs are close to dissolve, take a P1000 pipette to make single-cell suspension (set to 800 μL). If it is not fully dissolved, incubate for more time, till it gets dissolved. 9. Filter the plug suspension through a 70 μm strainer in a 50 mL Falcon tube. Rinse the strainer with 10 mL of PBS to assure that single-cell suspension goes in the tube (washing step). 10. Centrifuge the 50 mL Falcon tube at 200  g for 5 min. Remove supernatant very carefully, first with suction pump and then with P100 pipette. 11. Resuspend the pellet in 500 μL of PBS and transfer to 1 mL Eppendorf tube. Wash the 50 mL Falcon tube with 500 μL PBS and transfer to the 1 mL Eppendorf tube. Total volume will be 1 mL. 12. Centrifuge at 200  g for 5 min at 4
C (from now on everything should be on ice). 13. Dissolve pellet in 110 μL of PBS. 14. Transfer 10 μL from 110 μL to another 1 mL Eppendorf tube and add 200 μL of 2% formaldehyde. These tubes are named as beads, because PE-conjugated fluorescent counting beads will be added during FACS. Cover the tubes with parafilm and store at 4
C. 15. In the remaining 100 μL of cell suspension, add the suggested quantity of the dye of the staining kit for live/dead cells and incubate for 10 min in the dark at 4
C. 16. Add 1 μL of CD16/CD32 monoclonal antibody and 1.7 μL of 30% BSA in PBS (final concentration is 1% BSA), pipette gently with P100, and incubate at 4
C for 10 min in the dark. For multiple samples, make this mix in bulk and add in each tube. 17. Make a master mix of 1 μL from each antibody (Alexa Fluor 488-conjugated LYVE-1, PE-Cy7-conjugated CD31, APC-conjugated TER-119, APC-Cy7-conjugated CD45, and eFluor 450-conjugated CD102, 1 μL per sample). Incubate for 30 min on ice covered with aluminum foil with gentle shaking. 18. Wash with FACS buffer. Use at least 1 mL of FACS buffer per sample. Pipette gently and centrifuge at 1500 rpm for 5 min. Remove supernatant carefully. Repeat the wash twice. Total three washes are required. For first two washes, keep some

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Fig. 4 Gating strategy for characterization of vascular and lymphatic endothelial cells. Representative flow cytometry images from a highly vascularized Matrigel plug, treated with S1P and bFGF combination. The marked population in each graph corresponds to the cell population selected for next step analysis. The cells are first identified by a range of side scatter (SSC) and forward scatter (FSC), and then cell doublets are excluded (higher forward scatter), followed by dead-cell exclusion (higher indo-violet staining). This step provides the number of single alive cells of diverse origin that have invaded in the Matrigel plug. The next step leads to leukocyte exclusion (high CD45 staining) and red cell exclusion (high Ter-119 staining). The ICAM-2and CD31-positive cells are further separated in blood endothelial cells (low Lyve-1 expression) and lymphatic endothelial cells (high Lyve-1 expression)

liquid to avoid cell loss. For third wash, remove supernatant carefully as much as you can without having cell loss. 19. Add 500 μL of 2% formaldehyde, cover with parafilm and aluminum foil, and store at 4
C. Run the flow cytometry (FACS). 20. The gating strategy is schematically demonstrated at Fig. 4. Using this protocol, one can measure the number of total cells in the plug (general angiogenesis assessment), number of blood cells (demonstrating functional blood vessels), number of blood endothelial cells (demonstrating growth of blood vessels), and number of lymphatic endothelial cells (demonstrating growth of lymphatic vessels). 3.8 Directed In Vivo Angiogenesis Assay (DIVAA) (See Note 13)

In this modification, Matrigel is placed inside silicone tubes (angioreactors) that are then implanted in animals. Endothelial cells migrate inside the angioreactor and form vessels. This method prevents assay errors, such as Matrigel absorption from the mice, and is superior for accurate quantification of angiogenesis. Furthermore, it requires only small amounts of Matrigel and/or test

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substances, while up to four angioreactors can be implanted in each mouse, allowing fast and reproducible results with a small number of animals. More importantly, with proper optimization, it can be applied in different mouse strains. 1. Set angioreactors (see Notes 4 and 14) on ice (4
C), so that the open end is upside. 2. Gently fill the angioreactors with Matrigel that contains the tested substance(s) or the corresponding vehicle(s). 3. By using sterile forceps, turn the angioreactors upside down and place them in a sterile Eppendorf, to avoid the retraction of the Matrigel. 4. Place the angioreactors in closed Eppendorfs at 37
C up to 1 h to promote gel formation. 5. Anesthetize each mouse immediately before implantation, as described in Subheading 3.1, step 4. 6. Mice are shaved off hairs on the dorsal-lateral surface, approximately 1 cm above the hip socket. Then, using sterile scissors, make an incision after pinching back the skin and using sterile forceps, and insert the tube subcutaneously. Close the incision using sterile stitches. 7. Use 2–4 angioreactors for each mouse. 8. Matrigel incubation in the mice takes place for 9–15 days. 9. Euthanize mice (see Subheading 3.1, step 7). Exposure to CO2 levels greater than 70% for 5 min should be adequate. 10. Making small and careful incisions, remove the tubes carefully. Use a scalpel to carefully cut the blood vessels that have been formed into the tubes. 11. Remove the bottom cap of the silicone tube using sterile blade and push the gel out of the cylinder, inside a sterile Eppendorf tube. 12. Add 300 μL of digestion solution (see Note 12) for Matrigel digestion and gentle cell dissociation. Incubate at 37
C for 1–3 h. 13. Centrifuge digested Matrigel at 250  g for 5 min at RT to collect cell pellets and insoluble fractions, and discard supernatant. 14. Resuspend pellet in 500 μL of DMEM full medium to allow for cell-surface receptor recovery and incubate at 37oC for 1 h. 15. Centrifuge cells at 250  g for 10 min at RT to collect cell pellets. 16. Wash pellets three times with PBS.

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17. Add an appropriate fluorescently labeled lectin at an appropriate dilution (e.g., Griffonia simplicifolia lectin I, 1:200 in PBS) in every pellet and incubate overnight at 4
C. 18. Centrifuge cells at 250  g at RT and wash the pellet three times with PBS. 19. Resuspend pellet in 100 μL PBS and put in a plate for fluorometric detection. 20. Fluorescence is measured at excitation and emission filters that need optimization depending on the fluorochrome used, using a fluorescence plate reader. 21. The data are quantified in arbitrary units.

4

Notes 1. Growth factor-reduced Matrigel is used when stimulators of angiogenesis are to be studied. 2. The mice used in this assay must be of the C57BL/6 strain so that no effects related to the immune system interfere with the results. Other strains can be used only when the directed in vivo angiogenesis assay is employed that was initially developed in nude mice [5]. Optimizations are required for each strain. 3. Antibodies from many different suppliers can be used but conditions should be optimized in each case. 4. The angioreactors are usually included in commercially available kits but can also be prepared as follows: Section surgical grade silicone tubing of a 0.125 in. outside diameter, into 1 cm lengths and seal one end of the tubing with silicone adhesive. Rinse angioreactors in 70% ETOH followed by distilled water and sterilize using steam. 5. Matrigel is usually aliquoted and kept at 80
C. Before use, the aliquots are put in the refrigerator at 4
C overnight to defrost. Matrigel gels at 24–37
C in 30 min and once it is gelled it cannot redissolve. Pre-cooled pipette tips and tubes must be used. 6. The test substance(s) in the desired concentration is(are) mixed with the Matrigel prior to use and left overnight in the refrigerator to avoid bubbles. 7. During the procedure, when the animals are anesthetized, check for heart rate and respiratory rate every 15 min, if the procedure lasts longer than this. 8. The mice are shaved off hair on the dorsal-lateral surface of the mouse, approximately 1 cm above the hip socket.

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9. Mice are euthanized by exposure to CO2 levels greater than 70% for 5 min. 10. Small and careful incisions secure safe removal of the Matrigel plugs. 11. Other mounting solutions can be used as alternatives to Mowiol, as long as they are recommended for confocal microscope observation, optimized for better observation and stable fluorescence. 12. The quantity of the digestion solution required should be 1.5 mL per sample and normally, 1 extra sample should be calculated. 1 mL is used per sample and the other 0.5 mL is added should issues with dissolving exist. 13. The directed in vivo angiogenesis assay prevents some of the assay errors linked with the traditional Matrigel plug assay, such as Matrigel absorption from the mice. Furthermore, it requires only small amounts of Matrigel and/or test substances, while up to four silicone tubes can be implanted in every mouse, allowing for fast and reproducible results with a small number of animals. 14. Besides silicone tubes, other materials have been also used, such as Plexiglas ring/nylon net filter chambers [6].

Acknowledgments Pinelopi Kastana, Despoina Ntenekou and Stamatiki KatrakiPavlou are recipients of a PhD scholarship from the State Scholarship Foundation in Greece (IKY) (Operational Program “Human Resources Development – Education and Lifelong Learning,” Partnership Agreement PA 2014-2020). This research was supported in part by a Marie Curie Intra European Fellowship within the 7th European Community Framework Programme (ALTangioTARGET) and by the Intramural Research Program of the National Institute of Allergy and Infectious Diseases. All mice were bred and maintained under pathogen-free conditions at the University of Patras—Greece Animal Facility (EL13BIO04) or at American Association for the Accreditation of Laboratory Animal Care-accredited animal facilities at the NIAID or the TTUHSC School of Pharmacy and housed according to Directive 2010/63/ EU (http://eur-lex.europa.eu/legal-content/EN/TXT/? uri¼celex%3A32010L0063) or in accordance with the procedures outlined in the Guide for the Care and Use of Laboratory Animals under animal study proposals approved by the Veterinary Administration of Western Greece (approval # 118016/577/30-04-2014) or the NIAID or TTUHSC Animal Care and Use Committees, respectively. All mice were used between 6 and 13 weeks of age.

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References 1. Malinda KM (2009) In vivo Matrigel migration and angiogenesis assay. Methods Mol Biol 467:287–294. https://doi.org/10.1007/9781-59745-241-0_17 2. He´roult M, Bernard-Pierrot I, Delbe´ J, HammaKourbali Y, Katsoris P, Barritault D, Papadimitriou E, Plouet J, Courty J (2004) Heparin affin regulatory peptide binds to vascular endothelial growth factor (VEGF) and inhibits VEGF-induced angiogenesis. Oncogene 23:1745–1753. https://doi.org/10.1038/sj. onc.1206879 3. Adini A, Fainaru O, Udagawa T, Connor KM, Folkman J, D’Amato RJ (2009) Matrigel cytometry: a novel method for quantifying angiogenesis in vivo. J Immunol Methods 342:78–81. https://doi.org/10.1016/j.jim. 2008.11.016

4. Doc¸i CL, Mikelis CM, Lionakis MS, Molinolo AA, Gutkind JS (2015) Genetic identification of SEMA3F as an antilymphangiogenic metastasis suppressor gene in head and neck squamous carcinoma. Cancer Res 75:2937–2948. https://doi.org/10.1158/0008-5472.CAN14-3121 5. Guedez L, Rivera AM, Salloum R, Miller ML, Diegmueller JJ, Bungay PM, Stetler-Stevenson WG (2003) Quantitative assessment of angiogenic responses by the directed in vivo angiogenesis assay. Am J Pathol 162:1431–1439. https://doi.org/10.1016/S0002-9440(10) 64276-9 6. Kragh M, Hjarnaa PJ, Bramm E, Kristjansen PE, Rygaard J, Binderup L (2003) In vivo chamber angiogenesis assay: an optimized Matrigel plug assay for fast assessment of anti-angiogenic activity. Int J Oncol 22:305–311

Chapter 19 Exosomes from Cell Culture-Conditioned Medium: Isolation by Ultracentrifugation and Characterization Anurag Purushothaman Abstract Exosomes are small vesicles of endosomal origin secreted by most cell types. Recent studies have identified exosomes as important mediators of intercellular communication and as important source materials for many clinical applications, including minimal invasive disease diagnosis. Exosomes have been purified from in vitro cell culture supernatants by many different methods; however the most simple and reliable method involves purification by ultracentrifugation. This chapter describes a detailed protocol for isolating exosomes from cell culture-conditioned medium using ultracentrifugation and their characterization based upon size, number, and protein expression by several complementary methods such as transmission electron microscopy, nanoparticle tracking analysis, western blotting, and flow cytometry. Key words Exosomes, Ultracentrifugation, Conditioned medium, Western blot, Flow cytometry, Cell culture, Sucrose, Transmission electron microscopy

1

Introduction Exosomes are small membrane vesicles (30–120 nm diameter) that are secreted by most cells into the extracellular environment [1]. Exosomes are formed intracellularly and hence different from other microvesicles that are generated by direct budding from the plasma membrane [2]. Formation of exosomes starts by the inward budding of the limiting membrane of the endocytic compartments, leading to vesicle containing endosomes called multivesicular bodies [3]. These multivesicular bodies fuse with the plasma membrane to release their internal vesicles or exosomes into the extracellular space. Study on exosomes has recently attracted high interest because exosomes function in intercellular transfer of proteins, lipids, and diverse RNA molecules from exosome-producing cells to target cells [4]. Research over the past decade has well established that exosomes are important means for cells to function in several biological events like intercellular communication, metabolism, tumor metastasis, angiogenesis, and host immune response

Davide Vigetti and Achilleas D. Theocharis (eds.), The Extracellular Matrix: Methods and Protocols, Methods in Molecular Biology, vol. 1952, https://doi.org/10.1007/978-1-4939-9133-4_19, © Springer Science+Business Media, LLC, part of Springer Nature 2019

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[5]. In addition to their impact on cellular functions, exosomes also participate in remodeling the extracellular matrix using proteases and glycosidases on the surface of exosomes [6]. Exosomes have also drawn attention as attractive drug delivery system because of their increased stability in circulation, biocompatibility, and low toxicity [7]. Further, exosomes could serve as noninvasive biomarkers with diagnostic or prognostic roles, as they can be detected in all biofluids including blood, plasma, serum, cerebrospinal fluid, urine, saliva, and amnion fluid [4]. Exosomes are purified from nearly all mammalian cell types (such as endothelial cells, neural cells, plasma cells, muscle cells, stem cells) using multiple methods like ultracentrifugation, immunoaffinity isolation, polymer-based reagents, size-exclusion chromatography, and density gradient separation [8]. This chapter describes the simple and reliable ultracentrifugation method for the isolation of exosomes from conditioned media of cultures of cell lines. Further, before using these purified vesicles for any research purpose it is critical to ensure that the vesicles are exosomes and not any contaminating material from the conditioned medium. Different methods used for characterizing and accessing the purity of the isolated exosomes are described at the later part of this chapter. Ultracentrifugation is the most commonly used protocol for exosome purification. It involves several centrifugation and ultracentrifugation steps. Though ultracentrifugation protocol provides fairly pure exosomes, we also describe an extra purification step using sucrose cushion to remove more contaminants associated with exosomes. Because of the small size, exosomes can be visualized only with an electron microscope and therefore the evaluation of purity of exosomes and their characterization should be assessed using electron microscopy [9]. Different instruments such as Zetasizer Nano Range instruments and NanoSight are also available for determining the size and number of exosomes [10]. Further characterization of exosomes involves analysis of protein composition by western blot and flow cytometry [10]. A flowchart outlining the different steps involved in exosome isolation by ultracentrifugation and different methods used for exosome characterization is depicted in Fig. 1.

2

Materials

2.1 Exosome Isolation from Cell Culture-Conditioned Medium by Ultracentrifugation

1. Sterile phosphate-buffered saline (PBS) (137 mM NaCl, 2.7 mM KCl, 10 mM Na2HPO4, 1.8 mM KH2PO4, adjust the pH to 7.4 with HCl). 2. Cell culture complete medium with required nutrients (FBS, Lglutamine, antibiotics, 2-mercaptoethanol). 3. Serum-free culture medium (all the required nutrients except FBS).

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Fig. 1 Flowchart for exosome isolation by ultracentrifugation and their characterization. The first few steps in exosome isolation are designed to eliminate dead cells, large extracellular vesicles (EVs), and large cell debris. The ultracentrifugation step at 100,000  g pellet small vesicles corresponding to exosomes. The pellet is resuspended in PBS and ultracentrifuged one last time to eliminate contaminating proteins. Characterization of exosomes involves analysis of morphology, size/number, and protein composition

4. Ultracentrifuge and fixed-angle or swinging-bucket rotor. 5. Polycarbonate bottles or polyallomer tubes, appropriate for the ultracentrifuge rotor. 6. 0.22 μm Filter sterilization device. 7. 5 mL Syringe and 18G needle. 8. Tris/sucrose/D2O solution: 30 g Protease-free sucrose, 2.4 g Tris base, 50 mL D2O. Adjust pH to 7.4 with 10 N HCl drops and adjust volume to 100 mL with D2O. Sterilize by passing through a 0.22 μm filter and store up to 2 months at 4  C.

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2.2 Exosome Analysis 2.2.1 Transmission Electron Microscopy

1. Formvar-carbon-coated electron microscopy (EM) grids. 2. 2% Uranyl acetate: Weight 1 g of uranyl acetate and dissolve in 50 mL distilled water. Store up to 4 months at 4  C, in a 20 mL plastic syringe protected from light. Just before use, filter the amount needed of uranyl solution with a 0.22 μm filter. 3. 1% Glutaraldehyde: Dilute EM-grade glutaraldehyde fixatives (commercially available as 8%, 25%, or 70% aqueous solutions) in 0.1 M sodium phosphate buffer, pH 7.4, to appropriate dilution. Store up to 6 months at 20  C or up to 1 week at 4  C after thawing. 4. Sterile PBS. 5. Electron microscope. 6. Exosome pellet. 7. Whatman filter paper. 8. Parafilm.

2.2.2 Assessment of Exosome Size and Number

1. Zetasizer Nano range instruments (Malvern Nano-Zetasizer) or NanoSight NM20 instrument (NanoSight).

2.2.3 Flow Cytometry

1. MES buffer: Dissolve 0.489 g 2-(N-morpholino) ethanesulfonic acid (MES, free acid) in 80 mL ddH2O. Adjust to pH 6 with NaOH. Bring volume to 100 mL with ddH2O (final 0.025 M). Filter through 0.2 μm filter (do not autoclave this solution). Store unopened up to several months at 4  C. 2. Latex or magnetic beads. 3. Antibodies for coating beads (anti-CD63 or anti-MHC class II antibodies). 4. Ig isotype control antibody. 5. Fluorophore-coupled anti-exosomal protein antibody (e.g., anti-CD9, or anti-CD81). 6. 1 M Glycine in PBS. 7. 0.1% Bovine serum albumin (BSA) in PBS. 8. 1.5 mL Polypropylene microcentrifuge tubes. 9. Tube rotator. 10. Flow cytometer/fluorescence-activated cell sorter. 11. Exosome pellet.

2.2.4 Western Blot

1. Exosome pellet. 2. Whole-cell lysates of the same cell source as exosomes. 3. SDS sample buffer: 50 mM Tris–HCl, pH 6.8, 2% sodium dodecyl sulfate (SDS), 10% glycerol, 1% β-mercaptoethanol,

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12.5 mM EDTA, 0.02% bromophenol blue. Store at 4 to 25  C. 4. Tris-buffered saline (TBS): 50 mM Tris–HCl, pH 7.4, 150 mM NaCl. 5. Wash buffer: TBS with 0.05% Tween-20 (TBST). 6. Running buffer: 1 MES SDS running buffer-50 mM MES (2-[N-morpholino]ethanesulfonic acid), 50 mM Tris base, 1 mM EDTA, 0.1% SDS, pH 7.4. Store at 4  C to 25  C. 7. Blocking buffer: 5% Skim milk powder in TBST. 8. 4–12% Bis-Tris gels. 9. Electrophoresis equipment. 10. Transfer membrane (PVDF or nitrocellulose) and transfer unit. 11. BCA protein quantification reagents. 12. Antibodies to protein of interest. 13. Horseradish antibody.

peroxidase

(HRP)-conjugated

secondary

14. HRP substrate solution.

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Methods

3.1 Exosome Isolation from Cell Culture-Conditioned Medium by Ultracentrifugation 3.1.1 Preparation of Cell Culture-Conditioned Medium

For Adherent Cells

1. Grow cells in regular growth medium until 75% confluence. 2. Aspirate the medium and rinse cells three times with sterile PBS. 3. Add fresh serum-free culture medium (20 mL/15 cm dishes) (see Notes 1–3). 4. Incubate cells for 48 h at 37  C, 5% CO2 (see Note 4). 5. Collect the conditioned medium to 50 mL Falcon tube using a sterile pipet and proceed with exosome isolation (see Notes 5 and 6). For Non-adherent Cells

1. Grow cells in regular growth medium until they reach 70% of their maximum concentration. 2. Centrifuge cells at 300  g for 10 min, remove the medium by aspiration, and wash cells once in PBS. 3. Resuspend cells in fresh serum-free culture medium (see Note 7). 4. Incubate cells for 48 h at 37  C, 5% CO2 (see Note 4). 5. Collect the conditioned medium to 50 mL Falcon tube using a sterile pipet and proceed with exosome isolation (see Notes 5 and 6).

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3.1.2 Exosome Isolation by Ultracentrifugation

All centrifugations must be performed at 4  C. 1. Centrifuge the conditioned medium (from adherent or non-adherent cells) in the 50 mL Falcon tube at 300  g for 10 min (to pellet the cells). 2. Transfer the cleared conditioned medium to a new 50 mL Falcon tube leaving cell pellet behind (see Note 8). Centrifuge conditioned medium for 20 min at 2000  g, at 4  C (to pellet dead cells). 3. Transfer the supernatant to new polyallomer tubes or polycarbonate bottles appropriate for the ultracentrifugation rotor to be used and discard the pellet (see Note 9). 4. Centrifuge the supernatants for 30 min at 10,000  g, at 4  C (to pellet cell debris). 5. Transfer the supernatant to a fresh tube or bottle, leaving the pellet behind. Make sure not to contaminate the supernatant with the pellet (see Notes 9–11). 6. Centrifuge the supernatant for 70 min at 100,000  g, at 4  C (to pellet exosomes + contaminating proteins) (see Notes 12 and 13). 7. Remove the supernatant completely and discard (see Note 14). 8. Resuspend the pellet in each tube in 1 mL PBS and pool all the resuspended pellet from all the tubes containing materials from the same cell line to one single tube. Fill the tube with PBS (see Notes 15 and 16). 9. Centrifuge for 70 min at 100,000  g at 4  C. 10. Remove all of the supernatant with Pasteur pipet without touching the bottom of the tube where pellet is located (see Note 17). 11. Resuspend the exosome pellet in small volume (50–100 μL) of PBS (see Note 18). 12. Store exosomes up to 1 year at 80  C in 50 μL aliquots. Avoid repeated freezing and thawing (this can cause lysis of exosomes).

Purification of Exosomes on a Sucrose Cushion

Although the above ultracentrifugation protocol provides fairly pure exosomes, for some applications an extra purification step using a sucrose cushion is recommended. This step eliminates more contaminants associated with exosomes. 1. Resuspend the purified exosomes from step 12 in 25 mL PBS. 2. Make a cushion by loading 4 mL of Tris/sucrose/D2O to the bottom of SW8 tubes (tubes for SW8 swinging-bucket rotors for ultracentrifuge) (see Note 19).

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3. Gently add the diluted exosomes above the cushion without disturbing the interface and centrifuge for 75 min at 100,000  g, at 4  C. Using a 5 mL syringe fitted to an 18G needle, remove approximately 3.5 mL of the Tris/sucrose/ D2O cushion (which contains the exosomes) to a fresh ultracentrifuge tube and dilute to 70 mL with PBS. 4. Centrifuge for 70 min at 100,000  g, at 4  C, in a 45 Ti fixedangle rotor for ultracentrifuge. 5. Remove all of the supernatant with Pasteur pipet without touching the bottom of the tube where pellet is located (see Note 17). 6. Resuspend the exosome pellet in 50–100 μL PBS. 3.2 Exosome Analysis 3.2.1 Imaging of Exosomes by Transmission Electron Microscopy

1. Perform vacuum electrical discharge to make the formvarcarbon-coated EM grids temporarily hydrophilic. Place a drop of 10 μg exosomes in 5 μL PBS on a parafilm, invert grid on it, leave at room temperature for 1 min, and dry the drop with a Whatman filter paper. 2. Transfer the grid to a 50 μL drop of 1% glutaraldehyde for 5 min. 3. Transfer to a 100 μL drop of distilled water for 2 min. Repeat this seven times for a total of eight washes. 4. Add a 5 μL drop of 2% uranyl acetate on parafilm, invert grid on it, and leave for 5 min. 5. Remove excess liquid using a Whatman filter paper and air-dry the grid for 5 min. 6. Observe under the electron microscope at 80 kV. Images are captured under an Olympus digital camera (see Note 20).

3.2.2 Assessment of Particle Size and Number

Different methods are available for measuring the size and number of exosomes. These measurements are becoming mandatory for research involving exosomes. 1. The Zetasizer Nano range instruments (Malvern NanoZetasizer) measure particles from less than a nanometer size to several microns using dynamic light scattering and zeta potential. Add two different exosome concentrations of the same preparation (25 ng/μL and 50 ng/μL in PBS) while measuring the size and number (see Note 21). 2. Nanoparticle tracking analysis allows the determination of a size distribution profile of small particles with a diameter of 10–2000 nm in liquid suspension, using a NanoSight NM20 instrument (NanoSight). The sample should be diluted 10 ng/ mL and injected into a sterile syringe until the liquid reaches the tip of the nozzle (see Note 22).

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3.2.3 Flow Cytometry

1. Wash 25 μL of magnetic or latex beads with 100 μL MES buffer twice and dissolve the beads in 100 μL MES buffer. 2. Make the antibody mixture by mixing a volume equal to 12 μg of anti-CD63 or anti-MHC class II antibody with the same volume of MES buffer. Add the beads to antibody mixture and incubate overnight at 4  C, on a rotator wheel. 3. Microcentrifuge for 3 min at 4000 rpm (1503  g), remove the supernatant, resuspend the pellet in 1 mL PBS, add 110 μL of 1 M glycine (to make final concentration of 100 mM), and incubate at room temperature for 30 min. The purpose of this step is to saturate any remaining free binding sites on beads. 4. Microcentrifuge for 3 min at 4000 rpm (1503  g), remove the supernatant, resuspend beads in 400 μL PBS, microcentrifuge to wash beads twice with PBS, and dissolve the beads in 100 μL PBS containing 0.1% BSA. 5. Incubate 30 μg of purified exosomes, as measured by BCA protein assay (see Note 18) with 10 μL beads, overnight at 4  C rotating. 6. Microcentrifuge for 3 min at 4000 rpm (1503  g) at room temperature. Discard the supernatant and wash twice with PBS as in step 4. Resuspend beads in 50 μL PBS containing 0.1% BSA. 7. To 25 μL beads add 50 μL of fluorophore-coupled anti-exosomal protein antibody (e.g., anti-CD9, or anti-CD81) diluted in PBS containing 0.1% BSA and incubate for 30 min in the dark with gentle agitation. On another 25 μL of beads add an irrelevant isotype-matched control antibody for negative control staining (see Note 23). 8. Wash twice in PBS as in step 4. Resuspend beads in 200 μL PBS and analyze on a flow cytometry. If the fluorophore signal is weak with anti-exosomal protein antibody, try to increase the amount of exosomes.

3.2.4 Western Blot

To show that a given protein is specifically enriched in exosomes, it is important to compare on the same gel identical amount of protein from exosomes and whole-cell lysates of exosomeproducing cell. 1. To 10 μg of exosomes and total cell lysate in separate tubes, add PBS and 4 SDS sample buffer to make up the deserved volume. 2. Incubate at 95  C for 5 min. 3. Assemble the 4–12% gel in the electrophoresis unit and add the 1 MES SDS running buffer.

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4. Load the exosome sample and the cell lysate sample next to each other in the wells of the gel. 5. Turn on the power unit and run the gel at 150 V until the dye front reaches the bottom of the gel. 6. Assemble the gel and the membrane within the transfer unit and transfer the proteins to the membrane. 7. Remove the membrane from the transfer unit, place in a container with blocking buffer, and rock at room temperature for 1 h. 8. Pour off the blocking buffer and add primary antibody solution (anti-CD81, anti-CD9, anti-CD63, anti-clathrin, anti-flotillin1, anti-Alix), diluted in blocking buffer, onto the membrane and rock at 4  C, overnight. 9. Pour off the primary antibody solution, add wash buffer, and rock at room temperature for 5 min. Repeat this process two more times for a total of three washes. 10. Pour off the third wash and add the appropriate HRP-conjugated secondary antibody solution, diluted in blocking buffer. Incubate with rocking for 1 h at room temperature. 11. Pour off the secondary antibody solution, add wash buffer, and rock at room temperature for 5 min. Repeat this process two more times for a total of three washes. 12. Pour off the wash buffer, add prepared HRP substrate solution onto the membrane, and incubate for 1 min at room temperature. 13. Drain membrane of excess HRP substrate solution (do not let dry), wrap in plastic wrap, and expose to X-ray film.

4

Notes 1. For cancer cell lines fifteen to twenty 15 cm dishes should yield sufficient exosomes. The number of cell culture dishes should be optimized according to the cell type. 2. Use as many as cells necessary to produce at least 70 mL of conditioned medium for exosome purification. The yield of exosome purification increases with starting material so it is always better to start with larger volume of conditioned medium. 3. We prefer growing cells in serum-free media for preparing the conditioned medium for exosome isolation. If the cells do not survive in serum-free conditions, we prefer growing cells in FBS-containing medium prepared using FBS pre-depleted for

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exosomes. The following procedure can be followed for depleting exosomes from FBS; prepare medium with all required nutrients and 20% FBS. Centrifuge the medium overnight at 100,000  g (follow the instructions for centrifugations as explained in exosome purification section). Filter the sterile supernatant through a 0.22 μm filter. Make sure that the pellet does not move while pouring the supernatant to the filter. If it does, stop pouring and discard the remaining supernatant. Finally, dilute the filtered medium with medium containing all nutrients except FBS to reach the final FBS concentration required to make the exosome production media. For example, if 10% FBS-containing medium is required, dilute 1 volume of depleted medium (20% FBS) with one volume of FBS-free medium. Store the exosome-depleted diluted medium at 4  C for up to 4 weeks. 4. If the cells start dying in 48 h, collect the conditioned medium after 24 h. 5. Once the conditioned medium is collected, it is highly recommended to proceed with exosome isolation as soon as possible. Keeping the medium at 4  C for up to a week or at 80  C for months will lead to loss of exosomes and should be avoided. 6. It is always important to make sure that the cells are healthy and viable while collecting the conditioned medium for exosome purification. Otherwise it is possible that the product will be heavily contaminated with cell membrane fragments that are not exosomes. 7. For cancer cells such as myeloma cells, 0.7  106 cells/mL in 70 mL medium should yield sufficient exosomes. 8. Cell pellet at this step can be lysed by cell lysis buffer and the lysates can be stored at 80  C. This lysate can be used for western blot analysis along with exosomes at later time. 9. While transferring the supernatant make sure that none of the pellet is collected and contaminates the supernatant. Use a pipet rather than pouring off the supernatant. Hold the tube at an angle so that the pellet is always covered by the supernatant. Stop removing the supernatant when at least 500 μL of supernatant is still covering the pellet. 10. Before centrifuge mark one side of each centrifuge tube with a waterproof marker, at the bottom, and position the tube in the rotor with the mark facing up. In this way the mark will serve as a point to locate the pellet at the end of the centrifugation. In some cases, depending on the cell lines, there will probably not be a visible pellet at this step. If using swinging-bucket rotor the pellet will be located at the bottom of the tube. For fixedangle rotors, the pellet will be on the bottom side of the tube facing up (marked by the marker pen).

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11. For isolating exosomes in sterile conditions, use centrifuge tubes washed in 70% ethanol followed by sterile PBS and dried in hood. 12. Centrifugation time includes approximately 10 min for the centrifuge to reach 100,000  g and 60 min at the final speed. A longer time (up to 3 h) will not damage the exosomes. 13. For high-speed centrifugation at 100,000  g all tubes should be three-quarters full; if not use PBS to make the tubes threequarters full. 14. For removing the supernatant after the 100,000  g spin, for a fixed-angle rotor, pour off the supernatants rather than using a pipet. For swinging-bucket rotor, use a pipet to remove the supernatant and leave at least 500 μL of supernatant above the pellet. 15. In some cases there will not be a visible pellet at this point. If using swinging-bucket rotor flush the bottom of the tube. For fixed-angle rotors, resuspend by flushing up and down where the pellet is expected to be (bottom upper side of the tube, marked by the marker pen). 16. Minimizing the number of tubes for ultracentrifugation will increase the yield of exosomes because resuspension and pooling are less reliable and labor intensive when more tubes are used. 17. After the final spin to pellet down exosomes, remove all the supernatant using Pasteur pipet and leave behind approximately 100 μL of liquid above the exosome pellet. Use a micropipettor to remove the extra 100 μL without shaking the tube. 18. If the exosome pellet is used for western blot analysis, instead of resuspending in PBS the exosome pellet can be directly resuspended in 50 μL of 1 cell lysis RIPA buffer. Incubate on ice for 10 min and measure the protein content by BCA assay. 19. To remove the impurities in Tris/sucrose/D2O filter the solution first using a 0.22 μm filter, followed by a 0.1 μm filter, before using in ultracentrifugation. 20. Electron microscopy of exosomes reveals cup-shaped membrane vesicles. Often, smaller 10–20 nm lipid particles are also observed with the same preparations. 21. It is important to know the exact number of cells from which exosomes were isolated for normalization with the number of exosomes produced. 22. A concentration of 108–109 particles per mL works best for the accurate quantification of exosomes using NanoSight. An optimal dilution should present around 100 particles per field of view.

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23. For flow cytometry analysis of exosomes use a directly conjugated fluorescent antibody rather than antibodies requiring a secondary antibody step. Always perform a negative control staining using an irrelevant isotype-matched control antibody.

Acknowledgments This work was supported by a New Faculty Development Award from the Young Supporters Board of the UAB Comprehensive Cancer Center to AP. References 1. Purushothaman A, Bandari SK, Liu J, Mobley JA, Brown EE, Sanderson RD (2016) Fibronectin on the surface of myeloma cell-derived exosomes mediates exosome-cell interactions. J Biol Chem 291(4):1652–1663. https://doi. org/10.1074/jbc.M115.686295 2. Colombo M, Raposo G, Thery C (2014) Biogenesis, secretion, and intercellular interactions of exosomes and other extracellular vesicles. Annu Rev Cell Dev Biol 30:255–289. https://doi.org/10.1146/annurev-cellbio101512-122326 3. Thery C, Amigorena S, Raposo G, Clayton A (2006) Isolation and characterization of exosomes from cell culture supernatants and biological fluids. Curr Protoc Cell Biol Chapter 3:Unit 3.22. https://doi.org/10. 1002/0471143030.cb0322s30 4. Thery C, Zitvogel L, Amigorena S (2002) Exosomes: composition, biogenesis and function. Nat Rev Immunol 2(8):569–579. https://doi. org/10.1038/nri855 5. Vlassov AV, Magdaleno S, Setterquist R, Conrad R (2012) Exosomes: current knowledge of their composition, biological functions, and diagnostic and therapeutic potentials. Biochim Biophys Acta 1820(7):940–948. https://doi. org/10.1016/j.bbagen.2012.03.017 6. Sanderson RD, Bandari SK, Vlodavsky I (2017) Proteases and glycosidases on the surface of exosomes: newly discovered

mechanisms for extracellular remodeling. Matrix Biol. https://doi.org/10.1016/j. matbio.2017.10.007 7. Samanta S, Rajasingh S, Drosos N, Zhou Z, Dawn B, Rajasingh J (2018) Exosomes: new molecular targets of diseases. Acta Pharmacol Sin 39(4):501–513. https://doi.org/10. 1038/aps.2017.162 8. Greening DW, Xu R, Ji H, Tauro BJ, Simpson RJ (2015) A protocol for exosome isolation and characterization: evaluation of ultracentrifugation, density-gradient separation, and immunoaffinity capture methods. Methods Mol Biol 1295:179–209. https://doi.org/10. 1007/978-1-4939-2550-6_15 9. Thompson CA, Purushothaman A, Ramani VC, Vlodavsky I, Sanderson RD (2013) Heparanase regulates secretion, composition, and function of tumor cell-derived exosomes. J Biol Chem 288(14):10093–10099. https:// doi.org/10.1074/jbc.C112.444562 10. van der Pol E, Coumans FA, Grootemaat AE, Gardiner C, Sargent IL, Harrison P, Sturk A, van Leeuwen TG, Nieuwland R (2014) Particle size distribution of exosomes and microvesicles determined by transmission electron microscopy, flow cytometry, nanoparticle tracking analysis, and resistive pulse sensing. J Thromb Haemost 12(7):1182–1192. https://doi.org/ 10.1111/jth.12602

Chapter 20 Preparation and Characterization of Tissue Surrogates Rich in Extracellular Matrix Using the Principles of Macromolecular Crowding Adrian Djalali-Cuevas, Sergio Garnica-Galvez, Andrea Rampin, Diana Gaspar, Ioannis Skoufos, Athina Tzora, Nikitas Prassinos, Nikolaos Diakakis, and Dimitrios I. Zeugolis Abstract Tissue engineering by self-assembly allows for the fabrication of living tissue surrogates by taking advantage of the cell’s inherent ability to produce and deposit tissue-specific extracellular matrix. However, the long culture periods required to build a tissue substitute in conducive to phenotypic drift in vitro microenvironments result in phenotype and function losses. Although several biophysical microenvironmental modulators (e.g., surface topography, substrate stiffness, mechanical stimulation) have been used to address these issues, slow extracellular matrix deposition remains a limiting factor in clinical translation and commercialization of such therapies. Macromolecular crowding is an alternative in vitro microenvironment modulator that has been shown to accelerate extracellular matrix deposition by several orders of magnitude, thereby decreasing culture periods required for the development of an implantable device, while maintaining cell phenotype and function. Herein, we provide protocols for the production of tissue surrogates rich in extracellular matrix from human dermal fibroblasts, equine tenocytes, and equine adipose-derived stem cells using the principles of macromolecular crowding and the subsequent characterization thereof by means of immunofluorescent staining and complementary fluorescence intensity analysis. Key words Macromolecular crowding, Excluding volume effect, Cell therapies, Immunocytochemistry, Extracellular matrix

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Introduction Modern tissue engineering capitalizes the inherent capacity of cells to create native supramolecular assemblies. However, during in vitro expansion, deprived of their optimal tissue context, cells lose their phenotype and function. To overcome such issues, numerous in vitro microenvironments modulators, including

Adrian Djalali-Cuevas, Sergio Garnica-Galvez, and Andrea Rampin contributed equally to this work. Davide Vigetti and Achilleas D. Theocharis (eds.), The Extracellular Matrix: Methods and Protocols, Methods in Molecular Biology, vol. 1952, https://doi.org/10.1007/978-1-4939-9133-4_20, © Springer Science+Business Media, LLC, part of Springer Nature 2019

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surface topography, substrate elasticity, and mechanical loading, are at the forefront of scientific research and technological innovation [1]. The fabrication of native supramolecular assemblies, or tissue surrogates, is highly dependent on the rate of deposition of extracellular matrix (ECM) which, under traditional cell culture conditions, is extremely slow due to the very dilute culture conditions [2]. Macromolecular crowding (MMC) has been proposed as an efficient method to not only enhance ECM deposition, but also maintain their phenotype during in vitro culture [3]. MMC is a biophysical phenomenon that enhances thermodynamic activities and biological processes by several orders of magnitude [4, 5]. It is based on the addition of inert macromolecules to the culture media and it acts by recreating the highly confined or crowded in vivo environment [2]. Several molecules have been tested as macromolecular crowders, including dextran sulfate, Ficoll™, polyethylene glycol, polyvinylpyrrolidone, and carrageenan [6–10]. Among them, carrageenan, a highly sulfated polysaccharide, has been shown to induce maximum ECM deposition in permanently differentiated and stem cell cultures due to its higher polydispersity/ more effective excluding volume effect [3, 6, 11–16]. Immunofluorescent labeling techniques are a very valuable tool for in situ identification of a wide variety of biological molecules. This identification can be performed directly, by means of the use of a primary antibody against the molecule of interest labeled with a fluorochrome [17, 18], or indirectly by utilizing a primary antibody against the molecule of interest and a fluorescently labeled secondary antibody that targets specifically the primary antibody [19]. By using different combinations of primary and secondary antibodies conjugated with different fluorochromes, the spatial relations of two or three different molecules can be assessed simultaneously in one sample [20]. In addition, with the aid of a fluorescence microscope connected to a camera and publicly available software, a semiquantitative analysis can be performed in order to estimate the relative amounts of antigen that are present in different samples, thereby increasing the value of such techniques. Herein, we describe protocols for the fabrication of tissue surrogates rich in ECM from human dermal fibroblasts, equine tenocytes, and equine adipose-derived stem cells using the principles of MMC and the subsequent characterization thereof by means of immunofluorescent staining and complementary fluorescence intensity measurements.

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Materials All cell culture materials should be handled following aseptic technique. Human primary cells should be handled using Biosafety Level 2 practices and containment. Diligently follow all the waste disposal regulations of your country/institution when disposing any kind of waste material.

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1. Human normal adult dermal fibroblasts (hDFs), cryopreserved (ATCC). 2. Equine tenocytes (eTCs), cryopreserved. 3. Equine adipose-derived stem cells (eADSCs), cryopreserved. 4. Dulbecco’s modified Eagle’s medium, high glucose (DMEM). 5. Fetal bovine serum (FBS). 6. Penicillin-streptomycin solution 100 (PS). 7. Growth medium: Prepare growth medium by adding 10% FBS and 1% PS solution to the appropriated volume of DMEM. 8. Trypsin-EDTA solution. 9. Hanks’ balanced salt solution 10 (HBSS) without Ca2+ and Mg2+. 10. Double-distilled water, sterile. 11. Carrageenan powder, suitable for gel preparation. 12. 100 mM L-Ascorbic acid 2-phosphate solution: Prepare a stock solution of 100 mM L-ascorbic acid 2-phosphate in ddH2O and sterile filter with a syringe and a 0.2 μm syringe filter. Store it in frozen aliquots at 20  C and protect it from the light. 13. 0.2 μm Surfactant-free cellulose acetate sterile syringe filters. 14. 10 mL Luer-lock sterile syringes. 15. 75 cm2 Tissue culture flasks. 16. 48-Well tissue culture plates. 17. MMC growth medium: Weigh the appropriated amount of carrageenan powder in a 1.5 mL microcentrifuge tube in order to prepare 0.5 mL of medium per well to be treated, considering a concentration of carrageenan of 75 μg/mL (the concentration may need to be optimized for the cell population of interest). Always prepare 2 mL extra in order to compensate for pipetting errors. In order to disinfect the carrageenan, irradiate it with UV-C light for 15 min (see Note 1). Supplement growth medium with 1 μL of the 100 mM ascorbic acid solution per mL of medium in order to reach a concentration of 100 μM. This will be used as the control medium and will also be used to prepare crowding MMC growth medium by adding carrageenan. For the preparation of MMC growth medium, recover the disinfected carrageenan from the 1.5 mL microcentrifuge tube by suspending it in 1 mL of growth medium supplemented with ascorbic acid and transfer the volume to a tube with the remaining medium. Repeat this step at least twice in order to ensure the complete recovery of the carrageenan from the tube. For non-crowded control wells, keep aside the appropriated volume of growth medium supplemented with L-ascorbic acid 2-phosphate for this purpose (0.5 mL per well + 2 mL extra for compensating

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pipetting errors). For the solubilization of the carrageenan in the growth medium supplemented with ascorbic acid, incubate the tube in a thermostatic bath at 37  C for at least 30 min (see Note 2). 2.2 Immunofluorescent Characterization of the ECM

1. Phosphate-buffered saline (PBS): 137 mM Sodium chloride, 2.7 mM potassium chloride, and 10 mM phosphate buffer. Add 8 g of NaCl, 0.2 g of KCl, 1.44 g of Na2HPO4, and 0.24 g of KH2PO4 to 800 mL of dH2O. Adjust the pH to 7.4 with HCl before adding dH2O to 1 L. 2. Fixing solution: Paraformaldehyde (PFA) 2% (w/v) in PBS. Weight 0.2 g of PFA 95% powder and dissolve it by heating and stirring in a glass beaker containing 10 mL of 1 PBS (see Note 3). Allow dissolution for 1 h (see Note 4). Once the PFA has been completely dissolved, transfer the resulting fixation solution to a 50 mL centrifuge tube. Store at 4  C. Filter the solution through a 0.2 μm sterile syringe filter before use. 3. Blocking solution: 3% Bovine serum albumin (BSA) in PBS. Weight 0.3 g of bovine serum albumin and dissolve it in 10 mL of 1 PBS in a 15 mL centrifuge tube by vortexing. Keep it at 4  C for short-term or 20  C for long-term storage. 4. Primary antibodies (Abcam): Mouse monoclonal anti-collagen type I (ab90395), rabbit polyclonal anti-collagen type III (ab7778), rabbit polyclonal anti-collagen type IV (ab6586), rabbit polyclonal anti-collagen type V (ab7046), rabbit polyclonal anti-collagen type VI (ab6588), rabbit polyclonal antifibronectin (ab2413) (see Note 5). Prepare the dilutions of the primary antibodies in 1 PBS as indicated: mouse monoclonal anti-collagen type I (1:200), rabbit polyclonal anti-collagen type III (1:200), rabbit polyclonal anti-collagen type IV (1:200), rabbit polyclonal anti-collagen type V (1:200), rabbit polyclonal anti-collagen type VI (1:200), rabbit polyclonal anti-fibronectin (1:200). 5. Secondary antibodies (Thermo Fisher): Goat anti-rabbit conjugated to AlexaFluor®488 (A-32731), goat anti-mouse conjugated to AlexaFluor®555 (A-32727) (see Note 6). Prepare the dilutions of the secondary antibodies in 1 PBS as indicated: goat anti-mouse AlexaFluor®555 (1:500) for collagen type I immunofluorescent staining; goat anti-rabbit AlexaFluor®488 (1:500) for collagen types III, IV, V, VI; and fibronectin immunofluorescent staining. Protect them from exposure to light (see Note 7). 6. DAPI solution: Weight 1 mg/mL of DAPI and dissolve it in ddH2O to produce a 2000 solution. Dilute with methanol to obtain the 1 solution. 7. Mounting medium (Vectashield). 8. 8 mm Ø Round glass coverslips.

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1. Inverted fluorescence microscope, equipped with 20 objective, filters suitable for DAPI (Ex/Em: 358/461 nm), AlexaFluor®488 (Ex/Em: 495/519 nm), AlexaFluor®555 (Ex/Em: 555/565 nm), and a connected camera. 2. Q-Capture Pro 7 software for image acquisition or similar. 3. ImageJ (version: 1.51j8) software.

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Methods For the preparation of the tissue surrogates rich in ECM, routine cell culture practices should be applied.

3.1 Preparation of Tissue Surrogates Rich in ECM

1. Fill the appropriated number of 75 cm2 tissue culture flasks (T75) with 12 mL of growth medium. Equilibrate the flasks for 30 min in a cell culture incubator at 37  C and 5% CO2. 2. Thaw a vial of cryopreserved hDFs, eTCs, eADSCs, and low passage preferred (see Note 8) in a thermostatic bath at 37  C and seed the pre-equilibrated tissue culture flasks at 2500 cells/ cm2 for the hDFs or 5000 cells/cm2 for the eTCs and eADSCs. Incubate the cells overnight at 37  C and 5% CO2 under humidified atmosphere and replace the medium with 12 mL of fresh growth medium per T75 flask. Replace the growth medium every 3–4 days. 3. Once the cell cultures have reached approximately 85% confluency, discard the growth medium and wash the cell layers twice with 5 mL of 1 HBSS per T75 flask. Discard the HBSS, add 2 mL of trypsin-EDTA solution to each flask, and incubate for 5 min at 37  C. Assess cell detachment by microscopical examination and once all cells are detached from the flasks, add 4 mL of growth media, collect the cell suspension, and centrifuge it for 5 min at 400  g, 700  g, or 1200  g for the hDFs, eADSCs, and eTCs, respectively. Discard the supernatant and resuspend the cell pellet in 1 mL of growth medium per every cultured T75. Determine the cell concentration with the aid of a haemocytometer or similar. 4. To plate the desired number of wells in 48-well tissue culture plates, adjust cell concentration in order to seed 25,000 cells per cm2 for the hDFs and eTCs and 15,000 cells/cm2 for the eADSCs and ensure that each well has 0.5 mL of growth medium and incubate the plates at 37  C and 5% of CO2. In order to appreciate the effects of MMC on ECM deposition, at least three wells are used per immunofluorescence marker to be analyzed, and three wells without macromolecular crowder are used as control group. Plate the necessary number of wells per immunofluorescence assay in duplicate in order to have non-primary antibody controls in triplicate per condition.

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5. After 24 h of culture, examine the cells by microscopy and ensure that they are healthy and properly attached to the cell culture substrate before replacing the medium. 6. Replace the medium of the wells to be treated with MMC medium with 0.5 mL of the growth medium containing ascorbic acid and carrageenan or only ascorbic acid for control wells. Incubate the plates at 37  C and 5% CO2. 7. After 3 days of culture, the plates can be examined by microscopy and processed for immunofluorescent staining. 3.2 Immunofluorescent Characterization of the ECM

All the following steps must be performed at room temperature (unless otherwise stated). 1. At the end of every time point, aspirate the cell culture media of the corresponding plate and wash every well three times during 5 min with 500 μL of 1 sterile HBSS. Fix the samples with 150 μL/well of the filtered fixation solution pre-cooled at 4  C for 15 min (see Note 9). 2. Remove the fixation solution and wash the wells three times during 5 min with 250 μL of 1 PBS. Samples can be kept at 4  C in 1 PBS if immunofluorescent staining is not to be performed right away. 3. Add 150 μL per well of BSA blocking solution for 30 min to block unspecific binding sites for the primary antibody. 4. After the aforementioned 30 min, drain the blocking solution and incubate the samples with 75 μL per well of the corresponding primary antibody solution for 90 min. For negative control wells, add 75 μL of 1 PBS per well (see Note 10). 5. Remove the primary antibody solution and wash the wells three times during 5 min with 250 μL of 1 PBS (see Note 11). Incubate with 75 μL per well of the corresponding secondary antibody solution for 30 min. In this case, all the samples including the negative controls must be incubated with the corresponding secondary antibody solution (see Note 12). Protect the samples from light from here on. 6. Remove the secondary antibody solution and wash three times during 5 min with 250 μL of 1 PBS. 7. Remove the PBS and incubate the samples for 5 min with 75 μL per well of 1 DAPI solution in order to stain nuclei. 8. Remove the DAPI solution and wash three times during 5 min with 250 μL of 1 PBS. 9. Add 10 μL per well of VECTASHIELD® mounting media and place a coverslip in each well (see Note 13).

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Fig. 1 Immunofluorescent labeling of different extracellular matrix proteins in hDFs, eTCs, and eADSCs after 3 days in the presence (CR+) or absence (CR ) of macromolecular crowding (carrageenan). Macromolecular crowding increases deposition of matrix proteins in a cell type-dependent fashion

10. Analyze the samples by using ultraviolet excitation filter (e.g., Ex: 360–370 nm/Em: 420–460 nm) to visualize cell nuclei stained with DAPI, a blue excitation filter (e.g., Ex: 460–495 nm/Em: 510–550 nm) to visualize collagen types III, IV, V, and VI and fibronectin stained with AlexaFluor® 488, and finally a green excitation filter (e.g., Ex: 535–555 nm/Em: 570–625 nm) to visualize collagen type I stained with AlexaFluor®555 (Fig. 1). 3.3 Imaging and Quantification of Immunofluorescent Staining

Perform the steps relative to the image acquisition process in the dark, in order to prevent the loss of fluorescence intensity from the fluorophores as a consequence of light excitation. 1. Once the software has been launched, position the plate on the sample stage of the microscope and visualize a live preview from the camera input: be sure that the light path selector of the microscope is set on “camera” to view the image on screen. 2. It is pivotal to keep the same exposure time for all the samples examined for the same antigen (see Note 14). Adjust the

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exposure time for DAPI, and carefully adjust the focus on the cell layer. Select the correct filter to excite the fluorophore bound to the secondary antibody of interest, regulate the exposure time in order to avoid saturation, and take note of the longest exposure time void of saturation. Repeat these actions for all the samples, excluding the negative controls, and then set the lowest obtained exposure time value to examine all of the samples and controls. 3. Position the plate on the sample stage exposing the first sample to the light path, select the violet filter, and acquire a 20 magnified picture of the nuclei stained with DAPI. Without moving the sample, turn the filter block turret to the appropriated filter for the fluorophore bound to the secondary antibody of interest, regulate the focus, and acquire the image. Repeat this step five times in randomly selected fields of the well for each replicate of each condition. 4. Using the emission filter for the secondary antibody of interest, acquire five pictures from five randomly selected fields of the non-primary antibody-negative controls (see Note 15). Save all the acquired pictures in TIFF format before proceeding with the analysis of fluorescence intensity, which can be performed with the ImageJ software. 5. To proceed with the analysis, open the desired picture with the ImageJ software, open the “Analyze” menu, click on “Set measurement,” tick “Mean grey value” option, and then “OK.” Open again the “Analyze” menu and this time click on “Measure”: a results window will appear. Repeat this step for all the pictures of the replicate in examination and transfer the data to an Excel worksheet to further process them. Repeat this step for all the replicates of all the examined conditions, including the non-primary antibody-negative controls. 6. By using Excel average function, obtain the average of the mean gray values for each replicate, including negative controls, and then subtract the negative control mean value to each corresponding replicate to obtain their specific fluorescence intensity (see Note 16). 7. Calculate the average and standard deviation of each condition from the values of the corresponding replicates for further statistical analysis and graphical representations (Fig. 2). 8. To create a representative picture of the assay, open the desired image in ImageJ software together with the corresponding DAPI field. For both of the pictures, open the “Image” menu, select “Type,” and click on “32-bit.” Once all the images that need to be merged in a single picture have been turned to 32-bit format, open the “Image” menu, select “Colour,” and then “Merge channels”: match each picture

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with the desired channel, tick the “create composite” box, and click “Ok” to obtain the composite picture (see Note 17). Once the picture has been saved in the preferred format, it is possible to add a scale bar using the dedicated ImageJ tool (see Note 18).

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Notes 1. The efficacy of UV-C disinfection depends on many parameters, as are the intensity and wavelength of the UV radiation, the time of exposure, the distance from the source of

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irradiation, the presence of particles that can protect the microorganisms from UV, and a microorganism’s ability to withstand UV during its exposure. We currently disinfect the carrageenan with success inside of an open 1.5 mL microcentrifuge tube in vertical position with the UV lamps typically included in the biological safety cabinets used for cell culture, but more dedicated equipment can also be used with the same efficacy if similar conditions are met. 2. Once the carrageenan is suspended into the medium, if the tube is vortexed and observed through a light source, particles in suspension can be appreciated. Once the carrageenan is completely dissolved into the medium, no particles can be appreciated following the same observation procedure. It is very important to ensure the complete dissolution of the carrageenan into the medium as non-dissolved particles can deposit on the bottom of the plates and negatively affect the cell viability. 3. Avoid temperatures over 60  C (optimal are between 55 and 57  C), which result in methanol formation and reduction of the effective concentration of PFA. This can damage the cytoskeleton of the cells and increase the autofluorescence of the samples. 4. For safety reasons, make the fixation solution under a chemical hood and cover the glass beaker, with aluminum foil for example, to prevent the release of PFA toxic vapors and the evaporation of the PFA. Likewise, all the steps that involve the use of the fixation solution must be executed under a chemical hood due to the PFA toxicity. 5. To perform the localization of several molecules in the same sample by indirect immunofluorescent staining, it is required for the primary antibodies against the antigens of interest to be raised in different species in order to be able to be recognized by different specific secondary antibodies (e.g., the primary antibody anti-collagen type I has been raised in mouse and can be combined with any of the other primary antibodies raised in rabbit for their use in multicolor immunostaining). 6. For the selection of the secondary antibodies, several guidelines must be followed. First, the secondary antibody must be specific for the target species corresponding to the primary antibody (e.g., goat anti-mouse secondary antibodies for primary antibodies derived from mice and goat anti-rabbit secondary antibodies for primary antibodies derived from rabbit). Moreover, to avoid cross-reactivity with the non-intended primary antibody targets in multicolor immunostaining and increase specificity, secondary antibodies cross-adsorbed against IgGs from the nontarget species and sera from different species can

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be used (e.g., for the present study, we used goat anti-mouse secondary antibody cross-adsorbed against human serum and rabbit and goat IgGs, as well as goat anti-rabbit secondary antibody cross-adsorbed against mouse and goat IgGs). The selection of the fluorochromes conjugated with the secondary antibodies and the appropriated filter set is crucial to have the minimal signal overlapping and maximal specificity in case of detecting several molecules at the same time in multicolor immunofluorescent stainings. For this, it is very important to excite each fluorochrome with wavelengths as close as possible to its maximal excitation, avoiding at the same time to excite the other fluorochromes present in the sample. Also, for collecting the fluorescence of each fluorochrome specifically, the emission filter should match as close as possible the maximal emission wavelength of the corresponding fluorochrome, avoiding at the same time the emission of the other fluorochromes present in the sample. As an example, the filter set distribution and fluorochromes selected for the present study are shown in Fig. 3.

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7. Protect the samples from the light to avoid the loss of fluorescent signal due to photobleaching. The photobleaching effect (also termed as fading) causes a permanent photochemical alteration, by cleaving covalent bonds in the fluorochromes. These irreversible modifications make the fluorochrome unable to emit fluorescence and thereby decrease the signal in the samples. 8. (1) The hDFs that can be acquired from ATCC are generally in early passages (p0–p1). In order to expand them, before the preparation of the tissue surrogate, a cell density of 2500 cells per cm2 in growth medium was used. The subculture routine included 2  5 min washes with 5 mL of 1 HBSS, 2 mL of trypsin-EDTA solution for 5 min at 37  C, cell recovery with 4 mL of growth medium per each T75 flask, centrifugation for 5 min at 400  g, resuspension, counting, and plating at 2500 cells/cm2. Normally, the cells reach confluency after a week of culture, and medium is changed twice per week, using 12 mL per T75. For cell cryopreservation, a cell density of 500,000 cells/mL of 10% DMSO in FBS was used. (2) The eTCs used for this experiment were isolated by migration method from superficial digital flexor tendon of equine donors after surgical removal of the paratenon. Expansion in culture was performed using a seeding density of 5000 cells/cm2. Subculture was performed as for the case of the hDFs, except for the centrifugation of the cell suspension, which was performed at 1200  g, and for a centrifugation step which was added in the thawing procedure in order to remove DMSO traces before plating. For cell cryopreservation, a cell density of 1,000,000 cells per ml of 10% DMSO in FBS was used. (3) eADSCs were extracted by collagenase type I digestion from fat samples of the mane of four horses. eADSC subcultures were performed at 5000 cells/cm2 following the same procedure as for the hDFs, except for the centrifugation step at 700  g for 6 min. For cell cryopreservation, a cell density of 1.8–2  106 cells per mL of 10% DMSO in FBS was used. 9. Be extremely careful during fixation and washing steps to prevent the detachment of the cell layer. 10. Alternatively, incubation with the primary antibody solutions can be performed overnight at 4  C. 11. When working with a high number of experimental groups or samples, always proceed with the washes in a fractional way in order to avoid drying the samples. Drying may cause a background increase due to unspecific binding of the primary antibody to the sample. 12. In order to demonstrate the nonspecific binding of the secondary antibodies to the sample, the non-primary antibody control

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wells should be incubated with the corresponding secondary antibody solution. For multicolor immunostaining, these wells should be incubated with a solution containing the mixed secondary antibodies as applied to the sample wells. 13. Be extremely careful when placing the coverslips. Introduce them vertically in one edge of the well and leave them to fall slowly in order not to form bubbles and not to damage the cell layer. Do not press them forcefully once properly positioned, as this can cause the loss of the sample. 14. The selection of an optimal exposure time suitable for all the examined samples, defined as the maximal exposure time at which the sample with the highest fluorescence intensity does not show signs of saturation, will allow for the comparison of the collected data between the different samples. The acquisition of an excessively saturated image would result in an underestimation of fluorescence intensity. 15. Background fluorescence can result from several factors, such as nonspecific binding of the secondary antibodies, or autofluorescence from the culture plate, the cultured cells, and the collagen itself. The quantification of non-primary antibody controls fluorescence is therefore performed in order to estimate the amount of this nonspecific fluorescence, which will eventually be subtracted from the mean gray value measurements of the corresponding samples. 16. In order to proceed with further statistical analysis, values from each of the single replicates are needed. Therefore, subtract to the value of each replicate of the condition in analysis the mean value obtained from all the three replicates of the corresponding non-primary antibody control. 17. It is possible to merge more than two pictures, or channels, in this step, in order to appreciate the relative localization of the examined antigens. To reduce the background interference that can derive from nonspecific binding of the antibodies as well as from autofluorescence of several biological molecules, or even from the plastic substrate, the “Subtract background” ImageJ tool can be useful. To use it, right after the images have been turned to 32-bit format, open the “Process” menu, click on “Subtract background,” select the same rolling ball radius for every picture in the assay, confirm with “OK,” and then proceed with the merging of the channels. Choose a larger rolling ball radius to maximize the preservation of positive pixel intensity, or a smaller one to maximize background subtraction. 18. There are several options to add a scale bar with ImageJ. If the pixel/length ratio is known, it is possible to open the “Analyze” menu, select “Set scale,” type “1” in the “Pixels” box,

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type the corresponding distance in the “Known distance” box, then specify the unit of length, and confirm with “OK.” If the pixel/length ratio is not known, it is possible to use the “Straight” tool from ImageJ toolbar in order to trace a segmented line covering a known distance in the picture, then open the “Set scale” window, fill the “Known distance” and “unit of length” boxes, and then confirm with “OK.” A third option provides that a second picture, with the same pixel/ length ratio of the former and already containing its scale bar, is open and selected in ImageJ: in such case, it is only needed to tick the “Global” box in the “Set scale” window before clicking “OK” to apply the same scale to all the open pictures. To finally add the scale bar to the desired picture, all the listed options provide that the “Analyze” menu is open, in order to select “Tools > Scale bar,” personalize the scale bar, and confirm with “OK.”

Acknowledgments This work has been supported from the European Union, H2020 Research and Innovation Programme, ITN award, Tendon Therapy Train Project (grant agreement number: 676338); Science Foundation Ireland, Career Development Award Programme (grant agreement number: 15/CDA/3629); and Science Foundation Ireland and the European Regional Development Fund (grant agreement number: 13/RC/2073). The authors have no competing interests. References 1. Cigognini D et al (2013) Engineering in vitro microenvironments for cell based therapies and drug discovery. Drug Discov Today 18 (21):1099–1108 2. Chen C et al (2011) Applying macromolecular crowding to enhance extracellular matrix deposition and its remodeling in vitro for tissue engineering and cell-based therapies. Adv Drug Deliv Rev 63(4–5):277–290 3. Kumar P et al (2015) Macromolecularly crowded in vitro microenvironments accelerate the production of extracellular matrix-rich supramolecular assemblies. Sci Rep 5:8729 4. Minton AP (2001) The influence of macromolecular crowding and macromolecular confinement on biochemical reactions in physiological media. J Biol Chem 276(14):10577–10580 5. Zhou H-X, Rivas G, Minton AP (2008) Macromolecular crowding and confinement: biochemical, biophysical, and potential

physiological consequences. Annu Rev Biophys 37:375–397 6. Satyam A et al (2014) Macromolecular crowding meets tissue engineering by self-assembly: a paradigm shift in regenerative medicine. Adv Mater 26:3024–3034 7. Bateman JF, Golub SB (1990) Assessment of procollagen processing defects by fibroblasts cultured in the presence of dextran sulphate. Biochem J 267:573–577 8. Rashid R et al (2014) Novel use for polyvinylpyrrolidone as a macromolecular crowder for enhanced extracellular matrix deposition and cell proliferation. Tissue Eng Part C Methods 20(12):994–1002 9. Bateman JF et al (1986) Induction of procollagen processing in fibroblast cultures by neutral polymers. J Biol Chem 261(9):4198–4203 10. Lareu RR et al (2007) Collagen matrix deposition is dramatically enhanced in vitro when

Preparation of ECM-Rich Tissue Surrogates crowded with charged macromolecules: the biological relevance of the excluded volume effect. FEBS Lett 581(14):2709–2714 11. Satyam A et al (2016) Low, but not too low, oxygen tension and macromolecular crowding accelerate extracellular matrix deposition in human dermal fibroblast culture. Acta Biomater 44:221–231 12. Kumar P et al (2014) Macromolecular crowding: the next frontier in tissue engineering. Adv Sci Technol 96:1–8 13. Kumar P et al (2016) Low oxygen tension and macromolecular crowding accelerate extracellular matrix deposition in human corneal fibroblast culture. J Tissue Eng Regen Med 12:6–18 14. Daniela C et al (2016) Macromolecular crowding meets oxygen tension in human mesenchymal stem cell culture - a step closer to physiologically relevant in vitro organogenesis. Sci Rep 6:30746 15. Kumar P et al (2015) Accelerated development of supramolecular corneal stromal-like assemblies from corneal fibroblasts in the presence of

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macromolecular crowders. Tissue Eng Part C Methods 21:660–670 16. Chen B et al (2013) Macromolecular crowding effect on cartilaginous matrix production: a comparison of two-dimensional and threedimensional models. Tissue Eng Part C Methods 19(8):586–595 17. Coons AH, Creech HJ, Jones RN (1941) Immunological properties of an antibody containing a fluorescent group. Proc Soc Exp Biol Med 47:200–202 18. Coons AH, Kaplan MH (1950) Localization of antigen in tissue cells. J Exp Med 91:1–13 19. Coons AH, Leduc EH, Connoly JM (1955) A method for the histochemical demonstration of specific antibody and its application to a study of the hyperimmune rabbit. J Exp Med 102:49–59 20. Staines WA et al (1988) Three-color immunofluorescence histochemistry allowing triple labeling within a single section. J Histochemistry Cytochem 36:145–151

Chapter 21 Novel Approaches for Extracellular Matrix Targeting in Disease Treatment Nikolaos A. Afratis and Irit Sagi Abstract Extracellular matrix (ECM) macromolecules, apart from structural role for the surrounding tissue, have also been defined as crucial mediators in several cell mechanisms. The proteolytic and cross-linking cascades of ECM have fundamental importance in health and disease, which is increasingly becoming acknowledged. However, formidable challenges remain to identify the diverse and novel role of ECM molecules, especially with regard to their distinct biophysical, biochemical, and structural properties. Considering the heterogeneous, dynamic, and hierarchical nature of ECM, the characterization of 3D functional molecular view of ECM in atomic detail will be very useful for further ECM-related studies. Nowadays, the creation of a pioneer ECM multidisciplinary integrated platform in order to decipher ECM homeostasis is more possible than ever. The access to cutting-edge technologies, such as optical imaging and electron and atomic force microscopies, along with diffraction and X-ray-based spectroscopic methods can integrate spanning wide ranges of spatial and time resolutions. Subsequently, ECM image-guided site-directed proteomics can reveal molecular compositions in defined native and reconstituted ECM microenvironments. In addition, the use of highly selective ECM enzyme inhibitors enables the comparative molecular analyses within pre-classified remodeled ECM microenvironments. Mechanistic information which will be derived can be used to develop novel protein-based inhibitors for effective diagnostic and/or therapeutic modalities targeting ECM reactions within tissue microenvironment. Key words Extracellular matrix, ECM targeting, Matrix metalloproteases, Lysyl oxidases, Inflammation, Fibrosis, Cancer, Biological inhibitors

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Introduction A substantial part of tissue’s volume is the space between the cells, which is filled by an intricate network of macromolecules that constitute the ECM. A variety of proteins (e.g., collagen) and polysaccharides [e.g., glycosaminoglycans (GAGs), proteoglycans (PGs)] are secreted locally and assembled into the ECM organized meshwork in close association with the surface of their producer cell [1]. The common understanding of the functions of ECM macromolecules has evolved to viewing the ECM as a dynamic and versatile biomaterial, intricately regulating cell-cell connections,

Davide Vigetti and Achilleas D. Theocharis (eds.), The Extracellular Matrix: Methods and Protocols, Methods in Molecular Biology, vol. 1952, https://doi.org/10.1007/978-1-4939-9133-4_21, © Springer Science+Business Media, LLC, part of Springer Nature 2019

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interactions, and communications. This differs significantly from the historic perception of the ECM as a “static scaffold” that preserves tissue architecture. Thus, the functional unit in the body is not merely a cell, but a cell and the ECM which surround it. Accumulating evidence demonstrates that ECM orchestrates the cellular environment by maintaining metabolic equilibrium, strength, and elasticity as well as by affecting cell growth and differentiation [2]. The ECM is also a major molecular platform for the regulated action of proteolytic and cross-linking enzymes, such as matrix metalloproteases (MMPs) and lysyl oxidases (LOXs). Their known activities obtained from studies of normal and pathological tissues suggest that ECM proteolysis and ECM buildup may have striking roles in development as well as in disease states [3]. Thus, irreversible posttranslational modifications resulting from ECM enzymes will become increasingly important in the next decades of research and development as part of targeting many frontier problems in biology and medicine, while it will be critical to reach a detailed molecular understanding of the dynamic processes of ECM. However, masked by cellular components and without spatially or temporally classification, studying the heterogeneous ECM poses a great challenge [4]. The overall ECM morphology and structure are constantly undergoing remodeling where ECM components are deposited, degraded, or otherwise modified. Accordingly, biochemical processes occur over various timescales and are spatially dispersed within the tissue, creating a multi-complexed heterogeneous and dynamic molecular landscape. In accordance, study of the rapid postsynthetic modifications and degradation of ECM does not conform well to many powerful technologies, such as microarrays, proteomics, or genomics in order to decipher remodeling mechanisms with significant system-level and atomic detail molecular understanding. Thus, despite the overwhelming body of observations which have led to our present understanding of the ECM and the logical deductions that follow, key ECM mechanisms remain an enigma.

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The ECM Orchestrates Cellular, Tissue, and Organ Activities The ECM is created and oriented by the cells within it and takes two general forms: the interstitial matrix, which is a threedimensional gel of polysaccharides and fibrous proteins, and the basement membrane, which is a mesh-like sheet formed at the base of epithelial tissues [4, 5]. The role of ECM in cell adhesion and signaling [5] crystallized the notion that the functional unit in the body is not merely the cell, but the cell and the ECM that surround it. Studies demonstrated the following facts: cells do not grow unless properly anchored to the matrix, ECM may control generically erratic cells, and additionally cells can become corrupt in an

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abnormal extracellular environment [5–7]. In concurrence, studies of normal and pathological tissues have suggested that homeostasis of ECM macromolecules and its native fibers has striking effects in health and diseases [3]. Thus, irreversible ECM proteolytic processes are presumed to be well coordinated involving versatile dynamic and reciprocal dialogues, between ECM and cellular components. Changes in ECM macromolecule morphology, biomechanics, and general organization affect the microenvironment. For example, extensive tissue remodeling during cancer and inflammatory stages drew attention to enzymatic proteolysis of ECM and the resulted products and fragments (designated as matrikines) [7–9]. Defined enzymes, e.g., MMPs, a disintegrin and metalloproteinase with thrombospondin motifs (ADAMTS), plasmin, and cathepsin G (serine proteases), have been shown to degrade various yet selective ECM macromolecules, while ECM fragments and their receptors were shown in vivo to be directly involved in tissue remodeling during embryogenesis, tumor invasion [7], viral and bacterial infections, wound healing, and inflammation processes [9, 10]. The precise proteolysis of ECM bioactive molecules (cytokines, cell-adhesion molecules, and receptors) is redefining our views on the roles of several protease families in many physiological and pathological processes. Proteolysis of ECM by different proteolytic enzymes leads to distinct biological phenotypes, for example the highly structural homologous proteases, MMP-1 and MMP-13, and promotes differential degradation of ECM [11]. Even though both enzymes degrade collagen type I, MMP-13 has broader substrate specificity compared to MMP-1. MMP-13-mediated collagen type I fragments have greater length variety than MMP-1, suggesting that each enzyme produces intrapopulation heterogeneity. Moreover, ECM acts as a “reservoir” of several growth factors, cytokines, and other signaling molecules, and thus degradation of ECM releases these factors and regulates several cellular functions. It is demonstrated that ECM degradation by MMPs leads to exposure of adhesion sites (cadherins) and signaling molecules bound to ECM scaffold inducing signaling pathways, such as ERK1/2 [11]. In pulmonary fibrosis (PF) MMP-7 promotes the disease by cleavage of E-cadherin and activation of heparin-binding epidermal growth factor (HB-EGF) precursor to the active form [12, 13]. Moreover, MMP-8 is reported that cleaves IL-10 in murine model of PF and MMP-2 is linked to aberrant activation of Wnt/β-catenin signaling pathway [14, 15]. This underscores the idea that ECM proteolytic reaction mechanisms may represent master switches in the regulation of critical biological processes and cell behavior [3] and highlights that ECM-remodeling enzymes are crucial targets against many pathological conditions [16]. ECM consists of a multi-complexed molecular network with heterogeneous composition, organization, and dynamics: The

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non-coincidental complexity associated with ECM heterogeneity is based on unique physical, biochemical, and biomechanical properties embedded within its molecular network [17]. While fibrillar collagens and elastin contribute the major tensile strength and viscoelasticity of the tissue, fibronectin with laminin participates in building the matrix network, and PGs and GAGs function as modulators of the biological activities and as reservoirs of growth factors and cytokines. Due to this overwhelming complexity, complete molecular level understanding of how all these native tissue components are tightly organized in space and how this fine molecular organization altered during site-directed regulatory proteolysis is not fully available. Similarly, the hierarchical importance of ECM proteases, their specific activities, redundancy, temporal–spatial expression level and distribution, zymogen activation, protease turnover, and inhibition properties are unknown yet crucial issues in drug development [18–21]. Thus, the unmet need of the field is a new revolutionary experimental approach, which will enable the possibility to reveal the heterogeneous native ECM proteolysis and ECM cross-linking at fine molecular/atomic details with system-level understanding. Such experimental scheme will be invaluable to the rationalization of novel diagnostic and therapeutic agents targeting ECM pathophysiology. Owing to the intrinsic difficulties associated with characterization of the composition as well as the structural and biophysical properties of heterogeneous systems, the molecular/ atomic level understanding of ECM and its remodeling mechanisms remains elusive and fairly limited.

3 Native ECM Proteolytic Reactions in Health and Disease, Focusing on the Colon Organ The main proteolytic enzyme families, MMPs and ADAMs, are considered to be predominant proteases in ECM pathophysiological regulation. MMPs/ADAMs are a family of Zn+2-dependent ECM-degrading endopeptidases that share common functional domains and activation mechanisms and have the capacity to degrade all types of ECM proteins. Apart from playing a central role in ECM macromolecules turnover, these enzymes can proteolytically activate or degrade a variety of non-matrix substrates, including signaling molecules such as chemokines, cytokines, and growth factors. Accumulating data indicate that MMPs/ADAMs are the predominant proteases involved in the manifestation of various colon pathologies, which affect cellular function and migration (of inflammatory or tumor cells), mucosal ulceration, matrix deposition, and degradation, as well as the expression of signaling molecules [20]. Several MMPs are constitutively expressed and play

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a protective role through their effect on cellular homeostasis while others mediate tissue damage. As such, MMP-2, MMP-3, MMP-7, and MT1-MMP are constitutively expressed and regulate physiologic processes such as barrier function and mucosal defense. On the other hand, MMP-1, MMP-8, MMP-9, MMP-10, MMP-12, and MMP-13 are undetectable in normal intestine but their dysregulated expression during inflammation may play a role in cell adhesion, immune cell migration, and impaired wound healing. Despite the plenty of available data and general notion in the field, not much is known about the detailed action of MMPs/ ADAMs within the native colon tissues. Collagenases (MMP-1, -8, -13) are unregulated in inflammatory bowel disease (IBD) by myofibroblast macrophages, mesenchymal epithelial cells, and fibroblast. Together they degrade native collagens I–V and IX, gelatin, and proteoglycans. Gelatinases (MMP-2, -9) are unregulated in IBD by subepithelial and pericryptal fibroblast/myofibroblast, epithelial cells, and vascular endothelial and immune cells. Among their identified substrates denatured collagen, gelatin, laminin, elastin, fibronectin, aggrecan, vitronectin, and laminin can be found. Stromelysins (MMP-3, -10) are overexpressed during IBD by lamina propria, stromal cells, mononuclear macrophage cells, lymphocytes, and epithelial cells. Together they degrade gelatin and proteoglycans. Matrilysins (MMP-7, -12) are overexpressed in IBD by epithelial cells, macrophage-like cells of lamina propria, macrophages, and myofibroblasts. Their identified substrates are collagen, elastin, casein, heparin, and chondroitin sulfate. Finally, MT1-MMP is upregulated in myofibroblasts. It functions as MMP-2 activator and is responsible for scalping the colon ECM by degrading collagens I–III, gelatine, and fibronectin. Overall, regulated network activities of these ECM proteases maintain tissue homeostasis while their dysregulated activity during pathogenesis of IBD is responsible for tissue destruction and infiltration of inflammatory cells and microbes. Yet, the challenge remains to decipher their detailed zymogen activation processes and proteolytic reaction mechanisms toward their native ECM substrates. Thus, proteases may serve as master regulators by instructing pathological versus normal tissue homeostasis programs in the colon.

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LOXs in Health and Disease LOXs, a family comprising LOX and four LOX-like (LOXLs) enzymes, catalyze cross-linking of elastin and collagens. LOXs are ECM copper-dependent enzymes and the mechanism which catalyzes is between lysine and hydroxylysine residues that mediate the cross-linking of collagen, as well as lysine and lysine residues which mediate the cross-linking of elastin [22]. LOXs are involved in a

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spectrum of biological functions including cell migration, adhesion, and apoptosis and the expression of this molecule is regulated by growth factors and cytokines, such as hypoxia inducible factor-1, transforming growth factor β, and tumor necrosis factor α (TNFα) [23]. LOX elevation occurs in metastatic breast cancer, hepatic fibrosis, liver fibrosis, lung fibrosis, and cardiovascular diseases [24]. The catalytic activity of LOX family members is highly conserved and is mediated by the C-terminal domain which contains a copper-binding motif, lysine tyrosylquinone (LTQ) residue, and a cytokine receptor-like (CRL) domain [25]. On the other hand, the N-terminal region is differentiated among the members and it is important for the protein-protein interactions. Specifically, the LOX and LOXL1 prodomains are necessary for the secretion of the inactive proenzymes, which are then activated extracellularly. In contrast, LOXL2, LOXL3, and LOXL4 contain four scavenger receptor cysteine-rich (SRCR) domains instead of a pro-sequence and the SRCR domains are suggested that may be involved in protein-protein interactions [25]. Due to similarities in the domain arrangement, LOX and LOXL1 represent one LOX subfamily, whereas LOXL2–4 constitute another LOX subfamily. LOX expression has primarily been identified as a tumor suppressor, since the expression of LOX gene was found to inhibit the transforming activity of the H-Ras oncogene in NIH 3T3 fibroblasts [26]. LOX’s gene ability to inhibit Ras comes from propeptide domain of LOX protein. However, other studies showed data describing that reduced levels of LOX are associated with poor disease-free and overall survival [27]. These evidences raise the possibility that intracellular and extracellular LOXs have contrary functions. For example, in the case of invasive and metastatic breast cancer LOX is essential under hypoxia conditions and is suggested that hypoxia conditions increase the secreted active form of LOX, which enhances the mobility of the cells via enhancement of integrin activity and FAK activation [28]. Furthermore, in fibrotic diseases LOX promotes fibrogenesis of attenuated hepatic stellate cells and restricts fibrotic resolution in the case of liver fibrosis, whereas the bleomycin-induced pulmonary fibrosis leads to induction of epithelial to mesenchymal transition (EMT) in lung epithelium [29, 30]. Among the LOX family members, only LOXL1 levels were coincident with the appearance of cross-linked elastin in cirrhosis and co-localized with α-smooth muscle actin (α-SMA). Interestingly, the inhibition of LOXL1 in liver cirrhosis arrested expression of α-SMA, elastin, and collagen I [31]. LOXL2 remodels the tumor microenvironment and improves tumor cell survival and the overexpression was correlated with more aggressive breast cancer, lymph node metastasis, and reduced patient survival [32–34]. In addition, LOXL2 promotes hepatic progenitor cell commitment toward a cholangiocyte lineage and simultaneously reduces hepatocyte differentiation in liver fibrosis [35]. LOXL3

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secretion from myofiber enhances fibronectin oxidation, enables the enhancement of integrin activation, and ensures correct positioning and anchoring of myofibers along the myofiber stretch and integrin-mediated adhesion [36]. Finally, LOXL4 is upregulated in head and neck squamous cell carcinoma and promotes colorectal tumor metastasis [32, 37].

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Highly Potent Biological Therapeutic Agents Against Inflammation and Fibrosis Despite extensive efforts, the primary pathogenesis of major colon diseases’ processes remains elusive and current therapeutic approaches achieve suboptimal efficacy and safety profiles. As elaborated above, MMPs/ADAMs play crucial roles in disease initiation and progression. Thus, selective inhibition of native ECM proteolysis can be beneficial to control inflammation and tissue damage resulting from divergent inflammatory pathways. Yet, selective and effective drugs for inhibiting individual MMPs/ ADAMs have not yet been emerged, although massive research has been attributed over the past decade, along with multiple inhibitor class development and the outcome is summarized extensively in previous reviews [38–40]. The design of specific antagonists as drug candidates faces proven difficulties, since the most members of the family share high structural homology in their catalytic domains. Useful knowledge has been taken from the severe consequence of nonselective MMP inhibition by synthetic broad-spectrum inhibitors [41], elevated levels of natural MMP inhibitors, TIMPs, or highly specific ablation of individual protease [42–44]. Detailed mechanistic studies showed that structuraldynamic experimental approaches can be implemented to design potent and highly protein-based selective inhibitors targeting metalloproteinases in vivo. Based on these studies, such inhibitors against MMP-2, -7, -9, and -14 and TNFα-converting enzyme (TACE/ADAM17) were designed, mimicking endogenous-like inhibitor interactions in the form of active site-targeted functionblocking antibodies or allosteric functional-blocking recombinant proteins. Moreover, Sela-Passwell et al. have emphasized on how MMPs are tightly regulated as part of a “protease web” [21]. Interference with the “protease web” via a nonselective as well as highly selective MMP inhibition can induce systemic protease web-associated modulations (SPAM) [19]. A general and effective strategy unraveling key biomarkers is the application of different forms of proteomic analyses as global study of large numbers of proteins contained in a cell or an organism [45]. The recent development of functional profiling tools integrated into systems biology analysis is currently used to identify and quantify the dynamic functional interconnectivity of MMPs/ADAMs with other proteases, their inhibitors, and substrates in a cell or tissue at a

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particular time and per disease indication [46]. For example, the use of electron microscopy to analyze high-order structural and biophysical properties correlated to proteomics analysis detects major differences in the individual ECM components and diverse potential effects. This confirms the possibility of deciphering subtle structural and functional differences in selective protease interactions with their native substrates and focuses on identifying individual MMPs and their ligands as targets for disease therapies. However, current therapeutic approaches are characterized by lack of specificity without achieving the desired efficacy and safety profiles, which make the interpreting and validation of the results complicated [47]. Challenges related to designing selective and effective metalloprotease inhibitors are associated with the lack of detailed information about their specific biological pathways and biophysical action in native tissues [48]. Nevertheless, great effort has been done during the recent years to develop therapeutic antibodies against MMPs for targeting mainly fibrotic diseases and cancer. Implementing novel experimental strategy, our lab generated superselective functionalblocking antibodies against activated MMP-9, MMP-7, and MMP-14. For example, the selective blocking of MMP-2/MMP9, by a competitive inhibitory antibody (SDS3) targeting the active site of MMP-2/9, significantly attenuates inflammation in acute IBD mice models and reduces IL-6 in the inflamed tissue [49]. The immunization strategy for SDS3 was innovative, since an inorganic synthetic molecule, consisting of a Tris-imidazole zinc complex, Zn-tripod, which mimics the structure and chemical pattern of TIMP-binding mechanism toward the activated forms of MMP-2 and MMP-9, was used. The antibody was selected in order not to cross-react with free zinc ions or analogous metal-protein motif. Treatment with SDS3 has attenuated the severity of colitis in the DSS-induced murine model with considerable reduction of body weight loss, colon shortening, and diarrhea together with improvement of histopathological scoring. Apart from direct inhibition of active site of MMPs, other research groups have developed different approaches in which the antibodies are targeting not only the active site of the protease in the catalytic cleft, but also other regulatory loops and domains on the enzyme surface. An example of this strategy is the allosteric inhibitory antibodies, AB0041 and AB0046, which have been developed to target MMP-9 in IBD [50]. These allosteric inhibitory antibodies are highly selective and interact with the region close to the active site without interfering with the enzyme-binding cavity. Treatment of ulcerative colitis with MMP-9 inhibitors reveals significant alleviation of symptoms, such as body weight loss, diarrhea, and endoscopic disease, as well as reduction in histopathological findings and measured inflammatory factors, like TNFα [50].

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Based on the knowledge that ADAMs, as well as other proteases like MMPs, are expressed as zymogens and are activated upon proteolytic cleavage of their prodomain, our lab developed the prodomain of TACE/ADAM17 (TPD), as a potential therapeutic approach against TACE in rheumatoid arthritis and IBD. In vitro and in vivo data are describing that the blockage of TACE activity by rationally designed recombinant protein based on TACE-prodomain causes significant reduction of TNFα levels, inhibits effectively inflammatory response, and increases the survival rate in TPD trinitrobenzenesulfonic acid-induced colitis model [51]. TPD is highly selective toward TACE, but no effect was observed against other members of ADAM or MMP families [52]. Apart from ECM proteolysis, ECM stiffness is becoming an attractive area for drug designs especially in cancer and fibrotic diseases. For that purpose, specific antibodies targeting the LOXL2 have been developed, such as the allosteric monoclonal antibody, AB0024 which targets the domain between the third and the fourth SRCR regions. Despite the promising preclinical evaluation of this anti-LOXL2 antibody, the clinical trial had a disappointing outcome. The failure in idiopathic PF clinical study, due to lack of efficacy, directs the research interest to different indications such as liver fibrosis [35]. Recent article suggests that the anti-LOXL2 antibody, AB0024, leads to a deceleration in an advanced biliary and non-biliary liver fibrosis and promotes the reversal of fibrosis. Another antibody development approach, for more efficient inhibition of LOXL2, targeted its catalytic domain and named GS341. Treatment with this antibody in vitro and in vivo suggests the significance of LOXL2 in collagen fiber alignment and orientation, as well as the efficient regulation of this mechanism by this antibody. The crystal structure of LOXL2, which is now available, will provide useful information in order to design more efficient inhibitors against LOX(L) enzymes [53].

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Future Perspectives New strategies in order to study ECM homeostasis, molecular composition, and function within subcellular tissue microenvironments are well appreciated. The higher the resolution, the better and more complete the map will be. The desire is to directly observe native ECM molecules with fine molecular, near-atomic detail. Facing the above challenges, one of the main features of this novel scheme is the ability to perform image-guided site-specific biophysical, chemical, and biological studies with a system-level perspective, which will then be integrated to a complete view of the studied native ECM proteolysis environment. “Image-guided site-directed” proteomics will be applied to enable simultaneous

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Fig. 1 Schematic representation of ECM multidisciplinary integrated platform. Illustrated from left to right: native bio-samples are applied. A battery of biophysical, structural, and functional methodologies are exploited, through which bio-samples are distributed via a rational workflow and well-defined feeding loops that orchestrate back and forward flow of data, directing analyses. Moreover, the rationally designed feeding loops are enabling a breakthrough integration mode, bridging the gap between ECM macromolecule organization data and molecular action in atomic detail. The multifaceted data are rationally accumulated, analyzed, and managed via integrated algorithms, resulting in significant novel insights and outputs. These can be translated into a battery of therapeutic new agents, like protein-based inhibitors, diagnostics, medical devices, artificial tissues, and drug delivery strategies

analyses of many molecular species at a rationally selected site. Subsequently, protein-specific maps directly correlated with tissue architecture or morphology can be obtained (Fig. 1). Advancement in dynamic imaging and image reconstruction techniques, including differential interference contrast (DIC), confocal reflection microscopy, optical coherence tomography (OCT), two-photon microscopy, and second harmonic generation (SHG) imaging [54], now permits the nondestructive detection of cellECM/tissue interactions. These tools provide basic insights into physical arrangements of cells and ECM fiber and global insights into the mechanisms of tissue organization and remodeling. Fluorescent dyes are used to detect functions within cellular compartments and interfaces, such as tissue proteolysis, cell–cell interactions, and migration [55, 56]. These techniques have been

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implemented in studies of microenvironments within tissue models and have also been used to visualize live animals monitored by intravital microscopy [57, 58]. An important related area is the field of ECM biomechanics [59], which underlines how ECM reacts to various forms of force loads applied by cells residing in the matrix and the effect on cell behavior [60]. We suggest the development of biophysical structural-dynamic multimodality tools to study the tissue-remodeling proteins and enzymes at atomic detail [61, 62]. Utilization multimodality real-time kinetic X-raybased tools and atomic force microscopy imaging of single ECM enzyme molecules can show the balances within and between enzyme-substrate flexibility and structural kinetics [11, 63, 64]. In addition, in order to visualize the ECM and the surrounding tissue the development of airSEM™ microscopy provides the ability of high-resolution imaging, avoiding the time-consuming procedures of dehydration or coating for conventional scanning electron microscopy (SEM) [64]. airSEM™ is also easily integrated with any visualizing modality such as confocal, two-photon excitation, infrared, or atomic force microscopes. Moreover, the dualmetal staining, based on the combination of metal-containing histological and EM stains for tissue imaging such as ruthenium red and uranyl acetate, enhances the image contrast providing spatial and compositional mapping of ECM [64] (Fig. 1). Summarizing, it was suggested an entirely new and highly precise integrative research scheme, which is composed of (1) image-guided selective data acquisition strategy spanning micron to angstrom spatial resolutions, and days to subsecond time resolutions; [65] smart data integration and analysis tools enabling molecular holographic view of ECM; (2) research directed per specific native environment and pathological condition; and (3) rationalization of novel diagnostic and therapeutic agents inspired by ECM atomistic insights (Fig. 1). References 1. Theocharis AD, Karamanos NK (2017) Proteoglycans remodeling in cancer: underlying molecular mechanisms. Matrix Biol. https:// doi.org/10.1016/j.matbio.2017.10.008 2. Karamanos NK, Passi A (2014) Novel insights into matrix pathobiology regulatory mechanisms in health and disease. FEBS J 281 (22):4978–4979. https://doi.org/10.1111/ febs.13106 3. Lu P, Takai K, Weaver VM, Werb Z (2011) Extracellular matrix degradation and remodeling in development and disease. Cold Spring Harb Perspect Biol 3(12). https://doi.org/10. 1101/cshperspect.a005058

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INDEX A

F

Agarose gel electrophoresis ......... 59, 60, 63–65, 91–101 Aggrecan.............................. 2, 8, 10, 104–106, 108, 265 Angiogenesis......................................... v, vi, 2, 3, 5, 9–13, 15, 157–159, 166–167, 176–181, 211–231, 233 Atomic force microscopes........................................ v, 160, 165, 174, 175, 271 Autophagy .............................. v, vi, 2, 3, 12–14, 157–187 Avidin............................................................................. 134

Flow cytometry .......................................................vi, 104, 111–116, 226–228, 234, 235, 240, 244

G

Beclin 1 ...........................................................13, 158, 176 Biotin ........................ 112, 120, 123, 130, 131, 137–139

Gelatinases ............................. 9, 194, 202, 204, 207, 265 Gel filtration chromatography........................... 74, 76, 79 Gene expression ............................................. v, 22, 23, 28 Glycans.................................................... vi, 55, 56, 64, 81 Glycosaminoglycans (GAGs)..................................vi, 2, 7, 8, 10, 55–69, 71, 72, 75, 78, 79, 91, 108, 111, 118, 261, 264 Gouy-Chapman-Stern model ....................................... 144

C

H

Calcium ions......................................................... 144, 152 Charge screening effect ................................................ 144 Chelation ....................................................................... 152 Chondroitin sulfate (CS) .............................................2, 5, 6, 13, 56, 64, 66, 127, 144, 146, 147, 152 Collagens ..................................................... v, 2–4, 10–13, 21, 38, 39, 42, 44–52, 118, 158, 177, 183, 185, 194, 202, 203, 212, 215, 217, 218, 248, 251, 257, 261, 263, 265, 266, 269

Heparan sulfate (HS) ............ 2, 5, 6, 13, 15, 56, 64, 158 Heparin (Hep) ...................................................... 2, 5, 56, 57, 64, 212, 213, 265 High-performance liquid chromatography (HPLC)........................ vi, 59, 62, 68, 82–88, 129 Hyaluronan binding proteins (HABPs).................vi, 107, 108, 112, 113, 118, 120–122, 124 Hyaluronan (HA).....................................................v, vi, 2, 6–9, 21, 27, 81, 82, 85, 91–101, 103–109, 111–124, 127, 129, 131, 133, 138, 168 Hyaluronidases (HYALs) ............................................. v, vi, 6–8, 103–106, 108, 112, 113, 116, 127–140

B

D Decorin ................................................................ 4, 12, 13, 157, 160, 161, 167, 173, 181, 182 Digoxigenin...............................................................24, 26

E ELISA ...........................................................128–134, 137 Endorepellin (ER)......................................................3, 13, 157–161, 167, 172, 173, 176, 177, 182, 183 Endothelial cells .................................................... 1, 9, 13, 25, 158, 160, 163, 172, 173, 178, 185, 212, 216, 217, 219, 224–228, 234 Enteric nervous system (ENS) ............................ 117, 118 Excluding volume effect ............................................... 246 Exosomes............................................. v, vi, 5, 6, 233–244 Extracellular matrix (ECM).............................. v, vi, 1–15, 21–29, 33, 71, 72, 111, 117, 118, 143–155, 234, 245–255, 261–271

I Immunofluorescence .......................................... 104, 121, 123, 160, 165, 175, 176, 194, 225, 226, 246, 248–255 In situ gelatin zymography.................................. 193–198 In situ RT-PCR .........................................................21–29 Ion exchange chromatography............................... 59, 62, 63, 73, 75, 100

L LC3 .............................................................. 158, 176, 179

M Macromolecular crowding (MMC) ............... vi, 245–255 Marine invertebrates ............................................vi, 55–69

Davide Vigetti and Achilleas D. Theocharis (eds.), The Extracellular Matrix: Methods and Protocols, Methods in Molecular Biology, vol. 1952, https://doi.org/10.1007/978-1-4939-9133-4, © Springer Science+Business Media, LLC, part of Springer Nature 2019

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THE EXTRACELLULAR MATRIX: METHODS

278 Index

AND

PROTOCOLS

Matrigel ................................................................ 219–231 Matrigel plug........................................................ 219–231 Matrix metalloproteases (MMPs) ...................... 194, 198, 202, 203, 207, 262–264, 267, 268 Metabolic labeling.............................................. 72, 75, 77 Microvesicles .............................................................5, 233 Mitophagy ............................................... vi, 13, 160, 161, 167, 169, 181–182, 186 Molecular mass........................ vi, 7, 9, 91–101, 103, 131 Molecular mass distribution .....................................92, 95 Molecular weight ............................. 7, 92, 137, 139, 152 Myenteric ganglia............................................ vi, 118–124

N Nuclear magnetic resonance (NMR) ..................... 57, 60, 65–67, 69, 82

R Real-time gelatin zymography........................ vi, 201–209

S Scanning electron microscope (SEM) ................ v, 33–35, 37–39, 41, 43, 271 Spheroids .............................................................. 211–218 Sprouts........................................177, 212, 216, 225, 226 Starvation.............................................................. 160, 213 Substrate-electrophoresis .............................................. 128 Sulfated glycosaminoglycans ............................. 55–69, 91 Sulfated polysaccharides.............................. 55, 56, 65, 66

T Transmission electron microscope (TEM) ....... 33–37, 41

O

U

O-GlcNAc modification ................................................. 81 Osteogenesis imperfecta ................................................. 50

UDP-glucose................................................................... 82 UDP-glucuronic acid (UDP-GlcUA)......................83, 87 UDP-N-acetylglucosamine............................................. 82 Ultracentrifugation .............................................. 233–244

P Particle exclusion assay ................................... vi, 103–109 Patch-clamp ................................................. 146, 147, 153 Pericellular matrix ............................................ 2, 103–109 Polysaccharide purification ............................................. 63 Post-translational modifications .................. v, 45, 50, 262 Primary cell culture ........................vi, 118, 121, 124, 246 Proteases ....................................................... 9, 10, 45, 47, 93, 119, 131, 154, 160, 163, 167, 170, 193, 194, 198, 201–203, 207, 208, 234, 263–265, 267–269 Protein secretion ........................................................... 184 Proteoglycans (PGs) ........................................... v, vi, 2–5, 7, 8, 10, 12, 15, 38, 40, 61, 71–79, 104, 157, 158, 161–163, 169–173, 185, 261, 264, 265 Pulse-chase quantification .............................................. 50

V Voltage-gated ionic channels (VGICs) ...........6, 143, 145

W Western blotting.................................................... 79, 128, 160, 163, 164, 166, 173, 179, 185, 202, 234, 235, 237, 240–243

Z Zymography ................................................... vi, 128–131, 133–136, 139, 193–198, 201–209