Permeability Barrier: Methods and Protocols [2 ed.] 9781071616734, 1071616730

Citation preview

Methods in Molecular Biology 2367

Kursad Turksen Editor

Permeability Barrier Methods and Protocols Second Edition

METHODS

IN

MOLECULAR BIOLOGY

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

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

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

Permeability Barrier Methods and Protocols Second Edition

Edited by

Kursad Turksen Ottawa, ON, Canada

Editor Kursad Turksen Ottawa, ON, Canada

ISSN 1064-3745 ISSN 1940-6029 (electronic) Methods in Molecular Biology ISBN 978-1-0716-1672-7 ISBN 978-1-0716-1673-4 (eBook) https://doi.org/10.1007/978-1-0716-1673-4 © Springer Science+Business Media, LLC, part of Springer Nature 2021 This work is subject to copyright. All rights are reserved by the Publisher, whether the whole or part of the material is concerned, specifically the rights of translation, reprinting, reuse of illustrations, recitation, broadcasting, reproduction on microfilms or in any other physical way, and transmission or information storage and retrieval, electronic adaptation, computer software, or by similar or dissimilar methodology now known or hereafter developed. The use of general descriptive names, registered names, trademarks, service marks, etc. in this publication does not imply, even in the absence of a specific statement, that such names are exempt from the relevant protective laws and regulations and therefore free for general use. The publisher, the authors, and the editors are safe to assume that the advice and information in this book are believed to be true and accurate at the date of publication. Neither the publisher nor the authors or the editors give a warranty, expressed or implied, with respect to the material contained herein or for any errors or omissions that may have been made. The publisher remains neutral with regard to jurisdictional claims in published maps and institutional affiliations. Cover Illustration Caption: Artwork created by Kursad Turksen. This Humana imprint is published by the registered company Springer Science+Business Media, LLC part of Springer Nature. The registered company address is: 1 New York Plaza, New York, NY 10004, U.S.A.

Preface Over the last decade, significant new understanding has emerged on the components of permeability barriers and how they work in different tissues. However, our understanding of the formation, maintenance, regulation, and dynamics of permeability barriers is far from complete. Thus, the field remains a very active one, prompting me to put together a volume encompassing a series of protocols that reflect current investigations, the use of which will continue to help drive the area forward. Once again, the protocols gathered here are faithful to the mission statement of the Methods in Molecular Biology series: They are well-established and described in an easy to follow step-by-step fashion so as to be valuable for not only experts but novices in the stem cell field. That goal is achieved because of the generosity of the contributors who have carefully described their protocols in this volume, and I am grateful for their efforts. My thanks as well go to Dr. John Walker, the Series Editor of the Methods in Molecular Biology series, for giving me the opportunity to create this volume and for supporting me along the way. I am also grateful to Patrick Marton, the Managing Editor of Methods in Molecular Biology and the overall Springer Protocols collection, for his continuous support from idea to completion of this volume. A special thank you goes to Anna Rakovsky, Assistant Editor for Methods in Molecular Biology, for continuous support from beginning to end of this project. I would like to thank David C. Casey, Senior Editor of Methods in Molecular Biology, for his outstanding editorial work during the production of this volume. Finally, I would like to thank to production crew for putting together an outstanding volume. Ottawa, ON, Canada

Kursad Turksen

v

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

Use of Ussing Chambers to Measure Paracellular Permeability to Macromolecules in Mouse Intestine . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Doriane Aguanno, Ba´rbara Graziela Postal, Ve´ronique Carrie`re, and Sophie Thenet Rapid Evaluation of Intestinal Paracellular Permeability Using the Human Enterocytic-Like Caco-2/TC7 Cell Line . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Ba´rbara Graziela Postal, Doriane Aguanno, Sophie Thenet, and Ve´ronique Carrie`re Evaluation of Barrier Functions in Human iPSC-Derived Intestinal Epithelium . . . . . Shigeru Yamada and Yasunari Kanda Selective Regional Isolation of Brain Microvessels . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Fernanda Medina-Flores, Gabriela Hurtado-Alvarado, and Beatriz Go mezGonza´lez Estimating Brain Permeability Using In Vitro Blood-Brain Barrier Models . . . . . . . . . Saeideh Nozohouri, Behnam Noorani, Abraham Al-Ahmad, and Thomas J. Abbruscato In Vitro Human Blood-Brain Barrier Model for Drug Permeability Testing . . . . . . . . Ece Bayir and Aylin Sendemir Evaluation of Blood-Brain Barrier Integrity Using Vascular Permeability Markers: Evans Blue, Sodium Fluorescein, Albumin-Alexa Fluor Conjugates, and Horseradish Peroxidase . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Bulent Ahishali and Mehmet Kaya Blood-Brain Barrier (BBB) Permeability and Transport Measurement In Vitro and In Vivo . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Bingmei M. Fu, Zhen Zhao, and Donghui Zhu Assessment of Blood-Brain Barrier Permeability Using Miniaturized Fluorescence Microscopy in Freely Moving Rats . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Jeffrey L. Barr, G. Cristina Brailoiu, Ellen M. Unterwald, and Eugen Brailoiu Measurement of Lung Vessel and Epithelial Permeability In Vivo with Evans Blue . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Prestina Smith, Lauren A. Jeffers, and Michael Koval Measurement of Airway Epithelial Permeability: Methods and Protocols . . . . . . . . . . . ¨ calan ¨ ksel and Merve O Hasan Yu Vascular Permeability Assays In Vivo. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Mir S. Adil and Payaningal R. Somanath Endothelial Permeability Assays In Vitro . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Mir S. Adil and Payaningal R. Somanath

vii

v ix

1

13

27 37

47

73

87

105

123

137 149 165 177

viii

Contents

Mapping Receptor Antibody Endocytosis and Trafficking in Brain Endothelial Cells . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Mikkel R. Holst, Simone S. E. Nielsen, and Morten S. Nielsen An In Vitro Assay to Monitor Sertoli Cell Blood-Testis Barrier (BTB) Integrity . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Siwen Wu, Lingling Wang, Elizabeth I. Tang, Junlu Wang, and C. Yan Cheng Ussing Chamber Methods to Study the Esophageal Epithelial Barrier . . . . . . . . . . . . . Solange M. Abdulnour-Nakhoul and Nazih L. Nakhoul A Simple Method to Test Mechanical Strain on Epithelial Cell Monolayers Using a 3D-Printed Stretcher . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Amanda C. Daulagala, John Yost, Amirreza Yeganegi, William J. Richardson, Michael J. Yost, and Antonis Kourtidis Differentiating Between Tight Junction-Dependent and Tight Junction-Independent Intestinal Barrier Loss In Vivo. . . . . . . . . . . . . . . . . . . . . . . . . . . . Sandra D. Chanez-Paredes, Shabnam Abtahi, Wei-Ting Kuo, and Jerrold R. Turner Establishment of Intestinal Epithelial Cell Monolayers and Their Use in Calcium Switch Assay for Assessment of Intestinal Tight Junction Assembly . . . . . . . . . . . . . . . . Pawin Pongkorpsakol, Wilasinee Satianrapapong, Preedajit Wongkrasant, Peter R. Steinhagen, Nuttha Tuangkijkul, Nutthapoom Pathomthongtaweechai, and Chatchai Muanprasat A Method to Prepare a Bioprobe for Regulatory Science of the Drug Delivery System to the Brain: An Angulin Binder to Modulate Tricellular Tight Junction-Seal. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Keisuke Tachibana and Masuo Kondoh dSTORM Imaging and Analysis of Desmosome Architecture . . . . . . . . . . . . . . . . . . . . . Reena R. Beggs, William F. Dean, and Alexa L. Mattheyses Index . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

193

207

215

235

249

273

291 305 317

Contributors THOMAS J. ABBRUSCATO • Department of Pharmaceutical Sciences, School of Pharmacy, Texas Tech University Health Sciences Center, Amarillo, TX, USA; Center for Blood-Brain Barrier Research, School of Pharmacy, Texas Tech University Health Sciences Center, Amarillo, TX, USA SOLANGE M. ABDULNOUR-NAKHOUL • Departments of Medicine and Physiology, Tulane University School of Medicine, New Orleans, LA, USA SHABNAM ABTAHI • Laboratory of Mucosal Barrier Pathobiology, Department of Pathology, Brigham and Women’s Hospital and Harvard Medical School, Boston, MA, USA MIR S. ADIL • Clinical and Experimental Therapeutics, College of Pharmacy, University of Georgia, Augusta, GA, USA DORIANE AGUANNO • Centre de Recherche Saint-Antoine, INSERM UMRS 938, Sorbonne Universite´, INSERM, Paris, France; EPHE, PSL University, Paris, France BULENT AHISHALI • Department of Histology and Embryology, Koc¸ University School of Medicine, Istanbul, Turkey ABRAHAM AL-AHMAD • Department of Pharmaceutical Sciences, School of Pharmacy, Texas Tech University Health Sciences Center, Amarillo, TX, USA; Center for Blood-Brain Barrier Research, School of Pharmacy, Texas Tech University Health Sciences Center, Amarillo, TX, USA JEFFREY L. BARR • Center for Substance Abuse Research, Lewis Katz School of Medicine at Temple University, Philadelphia, PA, USA ECE BAYIR • Ege University Central Research Test and Analysis Laboratory Application and Research Center (EGE-MATAL), Izmir, Turkey REENA R. BEGGS • Department of Cell, Developmental, and Integrative Biology, University of Alabama at Birmingham, Birmingham, AL, USA EUGEN BRAILOIU • Center for Substance Abuse Research, Lewis Katz School of Medicine at Temple University, Philadelphia, PA, USA G. CRISTINA BRAILOIU • Department of Pharmaceutical Sciences, Jefferson College of Pharmacy, Thomas Jefferson University, Philadelphia, PA, USA VE´RONIQUE CARRIE`RE • Centre de Recherche Saint-Antoine, INSERM UMRS 938, Sorbonne Universite´, Paris, France; Paris Center for Microbiome Medicine (PaCeMM) FHU, APHP, Paris, Ile-de-France, France SANDRA D. CHANEZ-PAREDES • Laboratory of Mucosal Barrier Pathobiology, Department of Pathology, Brigham and Women’s Hospital and Harvard Medical School, Boston, MA, USA C. YAN CHENG • The First Affiliated Hospital, Wenzhou Medical University, Wenzhou, Zhejiang, China; The Mary M. Wohlford Laboratory for Male Contraceptive Research, Center for Biomedical Research, Population Council, New York, NY, USA AMANDA C. DAULAGALA • Department of Regenerative Medicine and Cell Biology, Medical University of South Carolina, Charleston, SC, USA WILLIAM F. DEAN • Department of Cell, Developmental, and Integrative Biology, University of Alabama at Birmingham, Birmingham, AL, USA BINGMEI M. FU • Department of Biomedical Engineering, The City College of the City University of New York, New York, NY, USA

ix

x

Contributors

BEATRIZ GO´MEZ-GONZA´LEZ • Area of Neurosciences, Department Biology of Reproduction, CBS, Mexico City, Mexico MIKKEL R. HOLST • Institute of Biomedicine, Aarhus University, Aarhus C, Denmark GABRIELA HURTADO-ALVARADO • Area of Neurosciences, Department Biology of Reproduction, CBS, Mexico City, Mexico LAUREN A. JEFFERS • Division of Pulmonary, Allergy, Critical Care and Sleep Medicine, Department of Medicine, Emory University School of Medicine, Atlanta, GA, USA YASUNARI KANDA • Division of Pharmacology, National Institute of Health Sciences (NIHS), Kawasaki, Japan MEHMET KAYA • Department of Physiology, Koc¸ University School of Medicine, Istanbul, Turkey; Koc¸ University Research Center for Translational Medicine, Istanbul, Turkey MASUO KONDOH • Graduate School of Pharmaceutical Sciences, Osaka University, Osaka, Japan ANTONIS KOURTIDIS • Department of Regenerative Medicine and Cell Biology, Medical University of South Carolina, Charleston, SC, USA MICHAEL KOVAL • Division of Pulmonary, Allergy, Critical Care and Sleep Medicine, Department of Medicine, Emory University School of Medicine, Atlanta, GA, USA; Department of Cell Biology, Emory University School of Medicine, Atlanta, GA, USA WEI-TING KUO • Laboratory of Mucosal Barrier Pathobiology, Department of Pathology, Brigham and Women’s Hospital and Harvard Medical School, Boston, MA, USA ALEXA L. MATTHEYSES • Department of Cell, Developmental, and Integrative Biology, University of Alabama at Birmingham, Birmingham, AL, USA FERNANDA MEDINA-FLORES • Area of Neurosciences, Department Biology of Reproduction, CBS, Mexico City, Mexico; Posgrado en Biologı´a Experimental, Universidad Autonoma Metropolitana, Unidad Iztapalapa, Mexico City, Mexico CHATCHAI MUANPRASAT • Chakri Naruebodindra Medical Institute, Faculty of Medicine Ramathibodi Hospital, Mahidol University, Samut Prakan, Thailand NAZIH L. NAKHOUL • Departments of Medicine and Physiology, Tulane University School of Medicine, New Orleans, LA, USA MORTEN S. NIELSEN • Institute of Biomedicine, Aarhus University, Aarhus C, Denmark SIMONE S. E. NIELSEN • Institute of Biomedicine, Aarhus University, Aarhus C, Denmark BEHNAM NOORANI • Department of Pharmaceutical Sciences, School of Pharmacy, Texas Tech University Health Sciences Center, Amarillo, TX, USA; Center for Blood-Brain Barrier Research, School of Pharmacy, Texas Tech University Health Sciences Center, Amarillo, TX, USA SAEIDEH NOZOHOURI • Department of Pharmaceutical Sciences, School of Pharmacy, Texas Tech University Health Sciences Center, Amarillo, TX, USA; Center for Blood-Brain Barrier Research, School of Pharmacy, Texas Tech University Health Sciences Center, Amarillo, TX, USA ¨ CALAN • Department of Pediatric Allergy and Immunology, Faculty of Medicine, MERVE O Celal Bayar University, Manisa, Turkey NUTTHAPOOM PATHOMTHONGTAWEECHAI • Chakri Naruebodindra Medical Institute, Faculty of Medicine Ramathibodi Hospital, Mahidol University, Samut Prakan, Thailand PAWIN PONGKORPSAKOL • Faculty of Medicine and Public Health, HRH Princess Chulabhorn College of Medical Science, Chulabhorn Royal Academy, Bangkok, Thailand BA´RBARA GRAZIELA POSTAL • Centre de Recherche sur l’Inflammation, INSERM UMR1149, Universite´ de Paris, Paris, France; Biology and Genetics of Bacterial Cell Wall Unit,

Contributors

xi

Pasteur Institute, Paris, France; Centre de Recherche Saint-Antoine, INSERM UMRS 938, Sorbonne Universite´, Paris, France WILLIAM J. RICHARDSON • Bioengineering Department, Clemson University, Clemson, SC, USA WILASINEE SATIANRAPAPONG • Section for Translational Medicine, Faculty of Medicine Ramathibodi Hospital, Mahidol University, Bangkok, Thailand AYLIN SENDEMIR • Department of Bioengineering, Ege University, Izmir, Turkey; Department of Biomedical Technologies, Ege University, Izmir, Turkey PRESTINA SMITH • Division of Pulmonary, Allergy, Critical Care and Sleep Medicine, Department of Medicine, Emory University School of Medicine, Atlanta, GA, USA PAYANINGAL R. SOMANATH • Clinical and Experimental Therapeutics, College of Pharmacy, University of Georgia, Augusta, GA, USA PETER R. STEINHAGEN • Department of Hepatology and Gastroenterology, Charite´ Medical School, Berlin, Germany KEISUKE TACHIBANA • Graduate School of Pharmaceutical Sciences, Osaka University, Osaka, Japan ELIZABETH I. TANG • The Mary M. Wohlford Laboratory for Male Contraceptive Research, Center for Biomedical Research, Population Council, New York, NY, USA SOPHIE THENET • Centre de Recherche Saint-Antoine, INSERM UMRS 938, Sorbonne Universite´, Paris, France; EPHE, PSL University, Paris, France; Paris Center for Microbiome Medicine (PaCeMM) FHU, AP-HP, Paris, Ile-de-France, France NUTTHA TUANGKIJKUL • Faculty of Medicine Ramathibodi Hospital, Mahidol University, Bangkok, Thailand JERROLD R. TURNER • Laboratory of Mucosal Barrier Pathobiology, Department of Pathology, Brigham and Women’s Hospital and Harvard Medical School, Boston, MA, USA ELLEN M. UNTERWALD • Center for Substance Abuse Research, Lewis Katz School of Medicine at Temple University, Philadelphia, PA, USA; Department of Pharmacology, Lewis Katz School of Medicine at Temple University, Philadelphia, PA, USA JUNLU WANG • The First Affiliated Hospital, Wenzhou Medical University, Wenzhou, Zhejiang, China LINGLING WANG • The First Affiliated Hospital, Wenzhou Medical University, Wenzhou, Zhejiang, China; The Mary M. Wohlford Laboratory for Male Contraceptive Research, Center for Biomedical Research, Population Council, New York, NY, USA PREEDAJIT WONGKRASANT • Department of Physiology, Faculty of Science, Mahidol University, Bangkok, Thailand SIWEN WU • The First Affiliated Hospital, Wenzhou Medical University, Wenzhou, Zhejiang, China; The Mary M. Wohlford Laboratory for Male Contraceptive Research, Center for Biomedical Research, Population Council, New York, NY, USA SHIGERU YAMADA • Division of Pharmacology, National Institute of Health Sciences (NIHS), Kawasaki, Japan; Pharmacological Evaluation Institute of Japan (PEIJ), Kawasaki, Japan AMIRREZA YEGANEGI • Bioengineering Department, Clemson University, Clemson, SC, USA JOHN YOST • Department of Surgery, Medical University of South Carolina, Charleston, SC, USA MICHAEL J. YOST • Department of Surgery, Medical University of South Carolina, Charleston, SC, USA

xii

Contributors

HASAN YU¨KSEL • Department of Pediatric Pulmonology, Faculty of Medicine, Celal Bayar University, Manisa, Turkey ZHEN ZHAO • Department of Physiology and Neuroscience, Zilkha Neurogenetic Institute, Keck School of Medicine, University of Southern California, Los Angeles, CA, USA DONGHUI ZHU • Department of Biomedical Engineering, Stony Brook University, Stony Brook, NY, USA

Methods in Molecular Biology (2021) 2367: 1–11 DOI 10.1007/7651_2021_367 © Springer Science+Business Media, LLC 2021 Published online: 18 March 2021

Use of Ussing Chambers to Measure Paracellular Permeability to Macromolecules in Mouse Intestine Doriane Aguanno, Ba´rbara Graziela Postal, Ve´ronique Carrie`re, and Sophie Thenet Abstract An increased intestinal permeability has been described in many diseases including inflammatory bowel disease and metabolic disorders, and a better understanding of the contribution of intestinal barrier impairment to pathogenesis is needed. In recent years, attention has been paid to the leak pathway, which is the route of paracellular transport allowing the diffusion of macromolecules through the tight junctions of the intestinal epithelial lining. While the passage of macromolecules by this pathway is very restricted under physiological conditions, its amplification is thought to promote an excessive immune activation in the intestinal mucosa. The Ussing chambers have been widely used to measure both active and passive transepithelial fluxes in intact tissues. In this chapter we present how this simple device can be used to measure paracellular permeability to macromolecules in the mouse intestine. We propose a detailed protocol and describe how to best exploit all the possibilities of this technique, correctly interpret the results, and avoid the main pitfalls. Key words FITC-dextran, Intestinal permeability, Leak pathway, Mouse intestine, Ussing chambers

1

Introduction Changes in the properties of the intestinal barrier have been the subject of growing interest in recent years as they have been associated with an increasing number of both digestive and extradigestive pathologies [1, 2]. In addition, the explosion of research on the intestinal microbiota has raised more and more questions requiring a better understanding of the reciprocal interactions between the microorganisms that are residing in the intestinal lumen and the host. At the center of these interactions, the intestinal barrier is a complex entity comprising the mucus layer covering the epithelium, antimicrobial peptides, and the underlying immune system, which cooperate together to protect the host against unwanted intrusion of harmful molecules or microorganisms and to prevent excessive immune activation. The epithelial cell monolayer and the intercellular junctions that tightly connect epithelial cells constitute the most crucial determinants of the physical barrier, acting as the gate-keeper between the external environment

1

2

Doriane Aguanno et al.

(i.e., the lumen content) and the remainder of the body. Tight junctions (TJ) form a selectively permeable barrier and constitute the rate-limiting step of paracellular transport, i.e., the passive flux of solutes through the space between cells. Tight junctions are made of highly dynamic and regulated molecular complexes forming an apical belt connected to the cell cytoskeleton [3, 4]. The intestinal epithelium is physiologically more permeable than other barriers in the body (such as the skin, the bladder, the distal segments of the renal tubules, the blood-brain barrier, etc.), making paracellular permeability through TJ an important contributor of transepithelial flux in this epithelium. A very fine control of paracellular permeability, decreasing from the small intestine to the colon and along the crypto-villous axis in the small intestine, is essential to maintain the function and homeostasis of the intestinal mucosa [4]. An intestinal hyperpermeability has been described in many pathological conditions [2] and the current paradigm proposes that the increased or uncontrolled passage of dietary or bacterial-derived molecules from the intestinal lumen can initiate (or at least contribute to) the inflammation observed in these pathologies, both in acute inflammatory states as in inflammatory bowel disease [2] and in the low-grade inflammation observed in metabolic diseases [5]. To better understand how gut barrier disruption can contribute to pathogenesis, there is a need to accurately measure variations of intestinal permeability in human samples and in animal experimental models. The Ussing chamber is a system that allows precise measurement of solutes flow through intact tissues. It was originally designed by the Danish Biologist Hans Ussing to measure active ionic fluxes in frog skin [6, 7]. Ussing chambers turned out to be also an accurate tool for assessing the passive flux of macromolecules between cells (i.e., paracellular permeability) in many epithelia, both with respect to ions and to macromolecules, whose control is ensured by distinct mechanisms [8] (see also chapter “Rapid evaluation of intestinal paracellular permeability using the human enterocytic-like Caco-2/TC7 cell line” by Postal et al. in this book). The measurement of ion permeability (ionic conductance) by measurement of transepithelial resistance (TEER) in Ussing chambers has been the subject of numerous reviews [6, 9, 10] and will not be discussed here. This chapter will focus on the use of Ussing chambers for the measurement of the permeability to macromolecules, which are probably more specifically involved in the initiation of local inflammatory response at the level of the intestinal mucosa and, ultimately, in other tissues, leading to systemic inflammation. The design of the Ussing chamber is illustrated in Fig. 1: the intestinal tissue sample is mounted at the interface of two hemichambers bathed with an oxygenated physiological buffer such as Ringer solution and maintained at 37  C. The mucosal (also

Measure of Intestinal Paracellular Permeability Using Ussing Chambers

3

Fig. 1 Ussing chambers system. (a) Photograph of a mounted and operating Ussing chamber system. 1: Mucosal (apical) Ussing chamber containing FITC-dextran 4 kDa; 2: Serosal (basolateral) Ussing chamber; 3: Tissue slider mounted with a sample; 4: Oxygenation tube; 5: heating block; 6: Plugs closing electrodes gaps. (b) Photographs of paired Ussing chambers (top) and tissue sliders (bottom). 7: Moieties of 2 mm ø aperture tissue sliders; 8: Moieties of 5 mm ø aperture tissue sliders with mounting pins. (c) Schematic representation of intestinal segment dissection and mounting onto tissue sliders. The intestinal tube (approximatively 1 cmlong segment) is opened by performing a longitudinal incision, leaving the mucosal surface facing up. The inner mucosal surface (luminal layer) corresponds to the apical side of the epithelium while the outward surface corresponds to the serosal side (basolateral compartment). The sample is gently transferred (mucosal side facing up) onto the tissue slider (the opaque one represented in gray) and centered on the aperture. The second moiety of the tissue slider (the transparent one represented in white) is then placed on top to block the sample. Both sliders are then placed between paired Ussing chambers. The chamber facing the mucosa corresponds to the luminal or apical pole of the epithelium while the chamber facing the serosal layer corresponds to the basolateral side (inner body compartment)

referred to as the apical or luminal) side of the tissue is facing one chamber half, whereas the serosal (also referred to as the basolateral) side is facing the other half-chamber. After a short period of equilibration, a known quantity of fluorescent tracer is added in the luminal chamber and its passage into the serosal chamber through the tissue is measured over time. The system is remarkably simple and it can be coupled with measure of the TEER [10]. Measuring paracellular permeability in mouse gut with Ussing chambers offers access to information that cannot be provided by overall permeability measurements such as the force-feeding of a tracer and its measurement in blood, another commonly used method to assess gut permeability [11]. The first one is that the permeability of distinct segments of the intestine can be measured specifically.

4

Doriane Aguanno et al.

Several genetic modifications give rise to a moderate hyperpermeability in all intestinal segments [11, 12] but in other experimental conditions, the permeability has been described to be differentially modulated along the digestive tract: for example, whereas the permeability was similarly increased in jejunum and ileum in mice fed a high fat diet, the hyperpermeability was much more marked in the jejunum than in the ileum in Ob/Ob mice [13]. A time-course study conducted in rats fed a high fat diet revealed that changes in paracellular permeability (assessed by flux of FITC-dextran 4 kDa) and transcellular permeability (assessed by flux of horseradish peroxidase) in the jejunum, ileum, cecum, and colon changed during the first weeks of the diet in a region-dependent manner [14]. A second asset of Ussing chamber is that permeability to tracers of different sizes can be assessed on the same sample. The importance of taking into account the permeability to macromolecules can be illustrated by the history of scientific work on occludin, a TJ-associated MARVEL protein (TAMP): measurement of ionic conductance and mannitol flux in the colon of occludin knockout mice showed no difference compared to wild-type mice and had temporarily led to the conclusion that occludin had no essential role in the barrier function [15]. A few years later, studies on cultured cell models using a variety of tracers ranging from mannitol to 70 kDa dextran demonstrated that occludin is in fact crucial for controlling permeability to macromolecules [16, 17], as does tricellulin, another TAMP protein, which is mainly localized at tricellular junctions [3, 18]. It has now become clear that the permeability to ions and to small molecules is controlled by proteins that differ in part from those involved in the control of the permeability to macromolecules [3, 8], with both common and distinct regulatory mechanisms. Much remains to be discovered in this area and thus, the variations in epithelial permeability under different pathophysiological conditions must consider potential impairment of the different paracellular routes.

2

Materials – Acrylic Ussing chambers placed on a heating block (EM-PMBLTY-6, World Precision Instruments, Fig. 1). The system must be connected to a circulating water bath. – Tissue sliders with a 2 mm ø aperture and matching the Ussing chambers (P2407, World Precision Instruments). See Note 1 and Fig. 1. – Plugs for closing electrodes holes (P2023, World Precision Instruments).

Measure of Intestinal Paracellular Permeability Using Ussing Chambers

5

– Oxygenation equipment: 95% O2 + 5% CO2 (carbogen) gas bottle, compatible manometer and control valve system (see Note 2). – Ringer buffer solution (115 mM NaCl, 25 mM NaHCO3, 1.2 mM MgCl2, 1.2 mM CaCl2, 2.4 mM K2HPO4, 0.4 mM KH2PO4) maintained at 37  C (see Note 3). – Phosphate Buffer Saline (PBS) solution containing 1 mM CaCl2 and 0.5 mM MgCl2. – 40 mg/mL stock solution of FITC-labeled dextran 4 kD, FD4 (see Note 4). – FITC-labeled dextran 4 kD standard concentrations (ranging from 10 μg/mL to 0.01 μg/mL). – Petri dishes. – Syringes and pipette tips for flushing. – Surgical vascular ball-tip and straight scissors, forceps. – 96-well plate and microplate fluorometer.

3

Methods

3.1 Setup of Ussing Chambers

1. Prior to the experiment, configure the circulating water bath on 42  C to maintain chambers at 37  C throughout the experiment and maintain Ringer solution and FD4 solution at 37  C. 2. Prepare the setup by placing the paired acrylic chambers onto the heating block (Fig. 1a, b). If no electrodes are used (for TEER measurements), the holes must be sealed by using appropriate tips.

3.2 Mouse Tissue Dissection and Mounting

1. After euthanasia, perform a midline incision to dissect the appropriate segment(s) to be analyzed, the small intestine and/or the colon. Depending on the number of replicates needed, the length of the sample will vary. A margin of half a centimeter should be added to flush the intestinal lumen without harming the mucosa (see Notes 5–7). 2. Using a syringe mounted a pipette tip, gently flush with cold PBS solution the intestinal lumen to remove feces, which can be collected. 3. In a petri dish, cut the sample into 1 cm-long segments. Each segment will be mounted in a separate slider (Fig. 1b, c). 4. Using surgical vascular ball-tip scissors, open the segment of intestinal tube by performing a longitudinal incision, leaving the mucosal surface facing up (Fig. 1c).

6

Doriane Aguanno et al.

5. With forceps, transfer the sample onto the tissue slider keeping the mucosa facing up (Fig. 1c). 6. Seal the slider, and place it on a paper soaked with PBS while waiting to be placed within the system. 7. Insert the slider with the mounted tissue between Ussing chambers and seal the system by firmly screwing each chamber. Take care to place the mucosal sides of all samples the same side (for example, the transparent part of the slider can be used as a visual clue for mucosal side). 8. Gently add 2 mL of Ringer solution per chamber and connect to the oxygenation system. 9. Leave each sample for equilibration at least 25 min (the duration of this equilibration phase will depend on the number of chambers to be mounted but should not exceed 1 h). Repeat to mount all chambers. On an indicative basis, to dissect and mount two replicates of tissue on two different sliders, 5 min are required and thus 30 min in total for 12 chambers. 3.3 FITC-Labeled Dextran 4 kDa (FD4) Addition and Flux Measurements

1. After equilibration, take and discard 100 μL of media from the mucosal chamber. Add 100 μL of FD4 stock solution (40 mg/ mL) to reach a final concentration of 2 mg/mL. Additional drugs or molecules of interest can be added to either the mucosal chamber or the serosal chamber at this stage or later (see Note 8 and Fig. 2). 2. Take 150 μL from each serosal chamber. These samples represent the basal fluorescence level at time-point 0. 3. Immediately read the fluorescence levels in 96-well plate using a fluorometer and standard concentrations to determine the tracer concentration in serosal samples. 4. Put the 150 μL samples back into each corresponding chamber to maintain unchanged media volume throughout the experiment. 5. Repeat steps 2–4 every 15 min (at least for 45 min) to measure the FD4 flux over time (see Note 9). 6. Mucosal media can be sampled at the end of the experiment to measure lactate dehydrogenase release, which is proportional to cell lysis, to assess tissue viability [19].

3.4 Disassembly and Cleaning of the System

1. Turn off the oxygenation system, oxygenation valves, and the heating system. 2. Remove Ussing chambers from the heating block, empty the remaining media, and soak them for half an hour in antibacterial detergent cleaning solution.

Measure of Intestinal Paracellular Permeability Using Ussing Chambers

A

B

Ex vivo measure of FD4 flux in the ileum

Linear regression of FD4 flux in the ileum 0.08

Control - duplicate 1 Control - duplicate 2 EGTA 5mM - duplicate 1 EGTA 5mM - duplicate 2 EGTA 15mM - duplicate 1 EGTA 15mM - duplicate 2

0.08 0.06

% FD4 in basal medium

% FD4 in basal medium

0.10

7

0.04 0.02 0.00 0

15

30

45

60

75

90

0.06

0.04

0.02

0.00

105

Control - duplicate 1 Control - duplicate 2 EGTA 5mM - duplicate 1 EGTA 5mM - duplicate 2 EGTA 15mM - duplicate 1 EGTA 15mM - duplicate 2

0

15

30

45

60

75

90

105

minutes

minutes

C

FD4 flux in ileum

slope (fold vs control)

8 6 4 2

M m 15 EG TA

TA EG

C

on

tr

5m

M

ol

0

Fig. 2 Example of permeability to FITC-Dextran 4 kDa (FD4) measured in mouse ileum with Ussing chambers. (a) FD4 passage measured over time in control or EGTA-treated ileum samples. EGTA is a calcium chelator known to rapidly disrupt intercellular junctions and thus leading to an increased paracellular permeability. EGTA was added in the mucosal chamber together with FD4 stock solution. Data are presented as the percentage of tracer in the serosal chamber over time. For each condition (control, EGTA 5 mM, EGTA 15 mM), two replicates of ileum samples were monitored (6 chambers). (b) Linear regression of the corresponding FD4 flux curves. The linear regression was applied on a linear part of FD4 kinetics. The first time-points were not included to take into account EGTA treatment effects. The obtained slope values for each curve were then analyzed as the output FD4 permeability parameter. (c) EGTA treatment effects on FD4 permeability in ileum. The experiment described in A was performed with two mice (12 chambers for a single run). Individual slope values of FD4 flux are presented in C for each condition (control, EGTA 5 mM, EGTA 15 mM) for two mice (represented by square and circle symbols) in duplicate. When further analyzing data, the replicates of each mouse are averaged

3. Remove the remaining tissues from the sliders and gently brush them with a toothbrush. Soak them for half an hour in antibacterial detergent cleaning solution. 4. Thoroughly rinse 3 times the chambers and slides in clean water (using filtered water for the last rinse) and leave them to dry. 5. Carefully flush the media reservoir and gas channels with air using a syringe to remove remaining water droplets.

8

3.5

Doriane Aguanno et al.

Data Analysis

1. Once all the concentrations of the fluorescent tracer are determined in each sample for the whole time-course, data can be analyzed as FD4 passage over time. Choose an appropriate part of the curve allowing to apply a linear regression (see Note 10) and calculate the slope of FD4 passage for each sample. This value is the final output parameter used to pool samples per condition and to compare conditions. 2. The above kinetics approach allows to identify damaged tissues through unexpected high values of fluorescence or slope changes. Other analysis methods are possible (see Note 10).

4

Notes 1. Depending on the available equipment and tissue samples, different types of tissue sliders with varying aperture diameters can be used (Fig. 1b). For mouse intestine, 2 mm ø aperture sliders are well suited (1 cm tissue samples). Five-mm ø aperture sliders with mounting pins allow a larger window of passage but this requires having larger tissue samples (1.5 cm), for example human surgical specimen. Smaller aperture sliders (0.8 mm ø) are adapted for small tissues such as biopsies. 2. The manometer must include a specific valve allowing a fine control of the gas pressure in order to maintain it low enough to generate a gentle bubbling in Ussing chambers. 3. Tissues can be also kept and processed for measurements in Ussing chambers in culture media such as Dulbecco’s Modified Eagle Medium (DMEM). Pay attention to glucose concentration, as it has been shown that a high glucose concentration leads to an opening of intercellular junctions and increase in paracellular permeability [7]. In any case, media should be phenol red-free to avoid noise autofluorescence. 4. The intestinal permeability can be assessed by the passage of different fluorescent dextrans with various sizes ranging from 4 kDa to 150 kDa, or smaller molecules such as FITC-labeled sulfonic acid 0.4 kDa [20]. The maximum size of a molecule so that it can cross the epithelium by the paracellular route remains debated. Its diameter could reach 100 Å (10 nm) [3, 17], which corresponds to the diameter of the “tube” formed at a tricellular junction [21]. It should be noted that many authors still consider that flux of tracers larger than 40 kDa evaluate exclusively the transcellular pathway; it is likely that, depending on the conditions, a mix of paracellular and transcellular pathways is assessed using such large tracers. It must also be kept in mind that epithelial damages generate an unrestricted, TJ-independent pathway [4]. In addition,

Measure of Intestinal Paracellular Permeability Using Ussing Chambers

9

fluorescent bacteria can be used to assess their passage across the intestinal barrier [12, 22]. Dextrans labeled with other fluorochromes such as TRITC-dextrans can also be used. The wavelengths of excitation and emission will depend on the fluorochrome and must be compatible with your microplate fluorometer. For FITC-Dextran, read the fluorescence with the following wavelengths parameters: 485 nm for excitation and 535 nm for emission. 5. For studies on mouse tissues, the seromuscular layer can be gently dissected and stripped from the mucosal layer [6], but this procedure could damage the tissue integrity and is not obligatory, as it has been shown in several studies [11, 12, 22]. To our knowledge, dissection of the submucosa and muscle plane is mandatorily performed for studies on human intestinal samples. 6. One can choose to compare intestinal permeability between the different gut segments (small intestine versus colon which feature distinct barrier function properties), or to analyze a single segment or even specific anatomical parts such as Peyer’s patches [12]. It is worth noting that the chosen segment of the digestive track will determine the number of possible replicates per mouse, the colon being much shorter than the jejunum for example. 7. Depending on the work hypothesis, the design of the experiment should consider the number of available Ussing chambers (6, 12, etc.), the different experimental conditions, and the number of samples that can be dissected from a single intestinal segment. If possible, two replicates per mouse and segment for a single condition should be analyzed for a more reliable analysis (Fig. 2c). As an example, with 12 available Ussing chambers and in the case of 3 experimental conditions, 2 mice per condition can be analyzed in a single run by mounting duplicate colon samples for each mouse. Thus, to analyze 8 animals from a same condition, four runs will be needed. 8. One of the advantages of Ussing chambers is the possibility to add treatments directly on the tissue ex vivo in a polarized manner (Fig. 2a–c). Compounds can be added either in the mucosal or serosal chamber. If added compounds lead to an increase in the tracer passage, the analysis should be performed on the linear section of the permeability curve (Fig. 2a–c). If the effect of the ex vivo treatment is observed within a timeframe compatible with the maintenance of the tissue viability, the two slopes of the tracer flow, before and after treatment, can be determined [20]. 9. If the tissues are kept in the system too long and their viability decline, the fluorescent tracer concentration will reach

10

Doriane Aguanno et al.

unusually high values. It is thus important to determine the optimal duration of the experiment in the course of optimization on test samples. As an example, in our conditions, the FD4 flux was still linear at 105 min (Fig. 2a). In the case of human biopsies or samples, the post-surgery delay is variable and must be taken into account. Keeping the sample into DMEM rather than a saline solution can help to preserve its viability [20]. If the human or mouse tissues have been impaired during dissection or mounting, aberrant values of fluorescence will be rapidly detected. 10. In a kinetic approach, FD4 detected in the serosal chamber can be plotted over time as either absolute values (quantity of tracer) or as percentage of the initial tracer quantity injected in the mucosal chamber [20]. The section of the curve of FD4 flux chosen to carry a linear regression analysis to obtain slope values should be the same for all replicates from a single experiment. Alternatively, the FD4 concentration measured at the end of the experiment can be expressed as flux (quantity/time/ area (depending of aperture surface of the sliders)) [11, 12, 23]. However, the slope values of the tracer passage measured over time seem to be more consistent between replicates than single time-point values.

Acknowledgments This work was supported by the Association Franc¸ois Aupetit (AFA); Institut National de la Sante´ et de la Recherche Me´dicale; Sorbonne Universite´; Ecole Pratique des Hautes Etudes; the Brazilian government’s Science Without Borders Program. DA was the recipient of a fellowship from CORDDIM Ile de France. B.G.P. received a doctoral fellowship (CNPq 207303/2014-2). References 1. Konig J, Wells J, Cani PD, Garcia-Rodenas CL, MacDonald T, Mercenier A, Whyte J, Troost F, Brummer RJ (2016) Human intestinal barrier function in health and disease. Clin Transl Gastroenterol 7(10):e196. https://doi.org/10. 1038/ctg.2016.54 2. Odenwald MA, Turner JR (2017) The intestinal epithelial barrier: a therapeutic target? Nat Rev Gastroenterol Hepatol 14(1):9–21. https://doi.org/10.1038/nrgastro.2016.169 3. Krug SM, Schulzke JD, Fromm M (2014) Tight junction, selective permeability, and related diseases. Semin Cell Dev Biol 36:166–176. https://doi.org/10.1016/j. semcdb.2014.09.002 4. Buckley A, Turner JR (2018) Cell biology of tight junction barrier regulation and mucosal

disease. Cold Spring Harb Perspect Biol 10(1): a029314. https://doi.org/10.1101/ cshperspect.a029314 5. Tilg H, Zmora N, Adolph TE, Elinav E (2020) The intestinal microbiota fuelling metabolic inflammation. Nat Rev Immunol 20 (1):40–54. https://doi.org/10.1038/ s41577-019-0198-4 6. Clarke LL (2009) A guide to Ussing chamber studies of mouse intestine. Am J Physiol Gastrointest Liver Physiol 296(6):G1151–G1166. https://doi.org/10.1152/ajpgi.90649.2008 7. Herrmann JR, Turner JR (2016) Beyond Ussing’s chambers: contemporary thoughts on integration of transepithelial transport. Am J Physiol Cell Physiol 310(6):C423–C431. https://doi.org/10.1152/ajpcell.00348.2015

Measure of Intestinal Paracellular Permeability Using Ussing Chambers 8. Shen L, Weber CR, Raleigh DR, Yu D, Turner JR (2011) Tight junction pore and leak pathways: a dynamic duo. Annu Rev Physiol 73:283–309. https://doi.org/10.1146/ annurev-physiol-012110-142150 9. Westerhout J, Wortelboer H, Verhoeckx K (2015) Ussing chamber. In: Verhoeckx K, Cotter P, Lopez-Exposito I, Kleiveland C, Lea T, Mackie A, Requena T, Swiatecka D, Wichers H (eds) The Impact of Food Bioactives on Health: in vitro and ex vivo models. Springer Open, Cham (CH), pp 263–273 10. Thomson A, Smart K, Somerville MS, Lauder SN, Appanna G, Horwood J, Sunder Raj L, Srivastava B, Durai D, Scurr MJ, Keita AV, Gallimore AM, Godkin A (2019) The Ussing chamber system for measuring intestinal permeability in health and disease. BMC Gastroenterol 19(1):98. https://doi.org/10.1186/ s12876-019-1002-4 11. Petit CS, Barreau F, Besnier L, Gandille P, Riveau B, Chateau D, Roy M, Berrebi D, Svrcek M, Cardot P, Rousset M, Clair C, Thenet S (2012) Requirement of cellular prion protein for intestinal barrier function and mislocalization in patients with inflammatory bowel disease. Gastroenterology 143 (1):122–132.e15. https://doi.org/10.1053/ j.gastro.2012.03.029 12. Barreau F, Meinzer U, Chareyre F, Berrebi D, Niwa-Kawakita M, Dussaillant M, Foligne B, Ollendorff V, Heyman M, Bonacorsi S, Lesuffleur T, Sterkers G, Giovannini M, Hugot JP (2007) CARD15/NOD2 is required for Peyer’s patches homeostasis in mice. PLoS One 2(6):e523. https://doi.org/ 10.1371/journal.pone.0000523 13. Johnson AM, Costanzo A, Gareau MG, Armando AM, Quehenberger O, Jameson JM, Olefsky JM (2015) High fat diet causes depletion of intestinal eosinophils associated with intestinal permeability. PLoS One 10(4): e0122195. https://doi.org/10.1371/journal. pone.0122195 14. Hamilton MK, Boudry G, Lemay DG, Raybould HE (2015) Changes in intestinal barrier function and gut microbiota in high-fat dietfed rats are dynamic and region dependent. Am J Physiol Gastrointest Liver Physiol 308(10): G840–G851. https://doi.org/10.1152/ajpgi. 00029.2015 15. Schulzke JD, Gitter AH, Mankertz J, Spiegel S, Seidler U, Amasheh S, Saitou M, Tsukita S, Fromm M (2005) Epithelial transport and barrier function in occludin-deficient mice. Biochim Biophys Acta 1669(1):34–42. https:// doi.org/10.1016/j.bbamem.2005.01.008 16. Al-Sadi R, Khatib K, Guo S, Ye D, Youssef M, Ma T (2011) Occludin regulates

11

macromolecule flux across the intestinal epithelial tight junction barrier. Am J Physiol Gastrointest Liver Physiol 300(6):G1054–G1064. https://doi.org/10.1152/ajpgi.00055.2011 17. Buschmann MM, Shen L, Rajapakse H, Raleigh DR, Wang Y, Wang Y, Lingaraju A, Zha J, Abbott E, McAuley EM, Breskin LA, Wu L, Anderson K, Turner JR, Weber CR (2013) Occludin OCEL-domain interactions are required for maintenance and regulation of the tight junction barrier to macromolecular flux. Mol Biol Cell 24(19):3056–3068. https://doi.org/10.1091/mbc.E12-09-0688 18. Krug SM, Amasheh S, Richter JF, Milatz S, Gunzel D, Westphal JK, Huber O, Schulzke JD, Fromm M (2009) Tricellulin forms a barrier to macromolecules in tricellular tight junctions without affecting ion permeability. Mol Biol Cell 20(16):3713–3724. https://doi. org/10.1091/mbc.E09-01-0080 19. Soderholm JD, Hedman L, Artursson P, Franzen L, Larsson J, Pantzar N, Permert J, Olaison G (1998) Integrity and metabolism of human ileal mucosa in vitro in the Ussing chamber. Acta Physiol Scand 162(1):47–56. https://doi.org/10.1046/j.1365-201X. 1998.0248f.x 20. Genser L, Aguanno D, Soula HA, Dong L, Trystram L, Assmann K, Salem JE, Vaillant JC, Oppert JM, Laugerette F, Michalski MC, Wind P, Rousset M, Brot-Laroche E, Leturque A, Clement K, Thenet S, Poitou C (2018) Increased jejunal permeability in human obesity is revealed by a lipid challenge and is linked to inflammation and type 2 diabetes. J Pathol 246(2):217–230. https://doi. org/10.1002/path.5134 21. Furuse M, Izumi Y, Oda Y, Higashi T, Iwamoto N (2014) Molecular organization of tricellular tight junctions. Tissue Barriers 2: e28960. https://doi.org/10.4161/tisb. 28960 22. Barreau F, Madre C, Meinzer U, Berrebi D, Dussaillant M, Merlin F, Eckmann L, Karin M, Sterkers G, Bonacorsi S, Lesuffleur T, Hugot JP (2010) Nod2 regulates the host response towards microflora by modulating T cell function and epithelial permeability in mouse Peyer’s patches. Gut 59(2):207–217. https:// doi.org/10.1136/gut.2008.171546 23. Laukoetter MG, Nava P, Lee WY, Severson EA, Capaldo CT, Babbin BA, Williams IR, Koval M, Peatman E, Campbell JA, Dermody TS, Nusrat A, Parkos CA (2007) JAM-A regulates permeability and inflammation in the intestine in vivo. J Exp Med 204 (13):3067–3076. https://doi.org/10.1084/ jem.20071416

Methods in Molecular Biology (2021) 2367: 13–26 DOI 10.1007/7651_2021_366 © Springer Science+Business Media, LLC 2021 Published online: 18 March 2021

Rapid Evaluation of Intestinal Paracellular Permeability Using the Human Enterocytic-Like Caco-2/TC7 Cell Line Ba´rbara Graziela Postal, Doriane Aguanno, Sophie Thenet, and Ve´ronique Carrie`re Abstract Paracellular permeability of the intestinal epithelium is a feature of the intestinal barrier, which plays an important role in the physiology of gut and the whole organism. Intestinal paracellular permeability is controlled by complex processes and is involved in the passage of ions and fluids (called pore pathway) and macromolecules (called leak pathway) through tight junctions, which seal the intercellular space. Impairment of intestinal paracellular permeability is associated with several diseases. The identification of a defect in intestinal paracellular permeability may help to understand the implication of gut barrier as a cause or a consequence in human pathology. Here we describe two complementary methods to evaluate alteration of paracellular permeability in cell culture, using the human intestinal cell line Caco-2 and its clone Caco-2/TC7. Key words Caco-2 cell line, Cell monolayer, Epithelium, FITC-dextran, Intestine, Paracellular permeability, Transepithelial electrical resistance

1

Introduction The intestinal epithelium is a cell monolayer mainly composed of enterocytes, which is at the interface between gut lumen and internal environment. The intestinal epithelium was defined in physiological situation as a “leaky” epithelium as compared to “tight” epithelia such as skin or urinary bladder [1–3]. It forms a selective barrier allowing the transfer of essential dietary nutrients, electrolytes, and water from the lumen into the circulation while preventing the entry of foreign antigens, microorganisms and their toxins [4, 5]. Two major routes are involved in this selective permeability: the transcellular and the paracellular pathways [4]. Transcellular pathway is involved in the absorption and transport of nutrients through processes predominantly mediated by specific transporters or channels located at the apical and basolateral membranes of intestinal epithelial cells [6]. Conversely, paracellular pathway corresponds to a passive but selective transport of only certain solutes and fluids through the space between adjacent cells [6]. Intestinal

13

14

Ba´rbara Graziela Postal et al.

PORE PATHWAY IONS & SMALL

LEAK PATHWAY MACROMOLECULES

MOLECULES

PARACELLULAR PERMEABILITY

TIGHT JUNCTIONS ADHERENS JUNCTIONS

DESMOSOMES

Fig. 1 Paracellular permeability is controlled by tight junctions and includes two distinct routes (the pore and the leak pathways), which implicate different tight junction proteins. As adherens junctions and desmosomes indirectly regulate tight junctions, their impairment may impact paracellular permeability

epithelial cells are connected by intercellular junction complexes localized all along the lateral membrane, but concentrated mainly at its upper part in what is called the “apical junctional complex.” The paracellular space is sealed at the level of tight junctions, which are the most apical cell-cell junctions and which specifically control the paracellular permeability [4, 5, 7, 8]. Paracellular permeability is divided in two pathways: the pore and the leak pathways (Fig. 1). The pore pathway is involved in the paracellular permeability to ions and small molecules and represents a high capacity flow depending on the size and charge of the molecules. The determination of transepithelial electrical resistance (TEER), which is inversely proportional to the flux of ions across the epithelium (ionic conductance), allows assessing the pore pathway. TEER value of the epithelium is the result of ionic fluxes through both the transcellular and paracellular routes but the contribution of the latter is predominant, especially in leaky epithelia such as intestine.

Epithelial Paracellular Permeability in Caco-2/TC7 Cells

15

As a consequence, the resistance at the tight junction level determines the overall resistance of the epithelium. The leak pathway is involved in the paracellular permeability to macromolecules of bacterial or food origin and displays a much lower capacity than the pore pathway in physiological conditions [9]. Monitoring the passage of large hydrophilic tracers using radiolabeled or fluorescent molecules such as dextrans reflects the leak pathway, which may vary depending on the size of the tracer used. It is important to note that distinct molecular mechanisms are involved in the regulation of leak and pore pathways [9]. Thus epithelial paracellular permeability to ions/small solutes (measured by TEER) versus macromolecules (measured by the passage of large molecules such as fluorescent-dextran) does not necessarily vary in parallel. Some experimental conditions can affect both pathways while other conditions may affect only one. Determination of paracellular permeability via methods analyzing pore and leak pathways is thus complementary. To study human intestinal paracellular permeability the challenge is to find an accurate model of intestinal epithelium close to normal human epithelium. Here we describe methods using the Caco-2/TC7 clone, which represents to date, with its parental cell line Caco-2, the most powerful in vitro model and the most used cell line to this purpose. The Caco-2 (Cancer colon-2) cell line was established in 1974 by Jorgen Fogh (Memorial Sloan-Kettering Cancer Center, New York, USA) from a human colon carcinoma taken from a 72-year-old man treated for his pathology with 5-fluorouacil and cyclophosphamide [10]. Among the several cell lines established from intestinal tumors by J. Fogh and others, some of them can be partially differentiated by adding synthetic or biological factors to the medium. However, one of them, the Caco-2 cell line was rather unique since it had the property to differentiate spontaneously upon reaching the confluence. The demonstration of Caco-2 cells differentiation was first published in 1983 [11] by Alain Zweibaum’s group (INSERM, Paris, France). They observed under standard culture conditions that Caco-2 cells spontaneously organize into a monolayer of polarized cells after reaching confluence (Fig. 2). Post-confluent Caco-2 cells display a well-defined brush border at their apical pole and high levels of intestinal hydrolases activities, which are, for some of them, close to the values observed in human normal small intestine [12, 13]. Since this first description of Caco-2 cell differentiation, this cell line has been the subject of numerous studies, which have shown that Caco-2 cells share a large number of properties with enterocytes of the normal human intestine. Thus post-confluent Caco-2 cells express a large number of proteins involved in several functions or metabolic pathways present in normal human enterocytes such as vitamin transport, nutrient absorption, and barrier function [14, 15]. Caco-2 cells have been widely used as a model of jejunum enterocytes for lipid absorption (952 PubMed hits) but it

16

Ba´rbara Graziela Postal et al.

Fig. 2 Post-confluent Caco-2/TC7 cells form a regular monolayer of wellpolarized cells with the nucleus in basal position and mature intercellular junctions as shown by immunofluorescence analysis of occludin, a tight junction marker (in green, panel A), E-cadherin, a marker of adherens junctions, (in green, panel B) and plakoglobin, a desmosomal protein (in green, panel C). Moreover, cells display a well-formed brush border as shown by actin staining (in magenta, panel B). Nuclei were stained with DAPI (in blue, all panels) (Figure adapted from [24])

is interesting to note that these cells express also some markers that are present in vivo in the human duodenum such as peptide transporters [16], and in the human ileum such as bile acid transporters [17]. Caco-2 cell monolayer cultured on plastic support is also characterized by the presence of domes, which result from apical to basal fluid transport, leading to uplift of the cell monolayer. Despite these characteristics, which bring them close to normal enterocytes, and due to their colonic and cancerous origin, these cells express also some markers of fetal colon (such as carcinoembryonic antigen) and display glucose metabolism characteristics of cancerous cells (high lactic acid production and accumulation of glycogen). In the early 1990s, the group of Ismael J Hidalgo (Absorption Systems LLC, Exton, USA) and the group of Per Artursson (Uppsala University, Sweden) identified the Caco-2

Epithelial Paracellular Permeability in Caco-2/TC7 Cells

17

cell line as a reliable and suitable cell model to study intestinal barrier function and in particular paracellular permeability [18, 19]. To date more than 700 publications reported data acquired with Caco-2 cells to study paracellular permeability. The Caco-2/TC7 cell line was established in the 1990s following the observation by the Zweibaum group of marked change in the expression of sucrase-isomaltase activity (a brush-border hydrolase) depending on the number of Caco-2 cell culture passages from the first establishment by J. Fogh (cells at late passages expressing more this enzyme than at early ones). They decided to establish several clones from an early passage (passage 29) and a late passage (passage 198) of the parental Caco-2 cell line. By dilution cloning, Monique Rousset (from the Zweibaum lab) isolated 26 cellular clones and compared their differentiation characteristics [20]. One of them, the TC7 clone (“T” for passage “Tardif” (the French translation of “late”) and C7 for the position of the well in the 96-well plate used for cloning) was the one that displayed the higher expression of several differentiation markers (brush-border hydrolases, sugar transporters) as compared to the others [20, 21]. The Caco-2/TC7 cells displayed some differences with the parental Caco-2 cells, e.g., they do not form domes when cultured on plastic support. Moreover, Caco-2/TC7 cells represent a more homogeneous cell population than the parental cell line, giving better reproducibility of results. An inter-laboratory study showed that Caco-2/TC7 cells exhibit higher alkaline phosphatase activity compared to parental Caco-2 cell line [21]. Caco-2 and Caco-2/TC7 cell lines display TEER values [21–25], which approximate those measured ex vivo in the human intestine while remaining higher [24, 26]. To date, among the existing intestinal cell lines, the Caco-2 cell line and particularly the Caco-2/TC7 clone represent the cell model that exhibit the morphological and functional differentiation closest to normal human enterocytes.

2 2.1

Materials Cell Culture

1. Caco-2/TC7 cells (see Note 1). 2. Complete culture medium: high-glucose (4.5 g/L) Dulbecco’s modified Eagle’s medium (DMEM) Glutamax supplemented with 20% heat-inactivated fetal bovine serum (FBS), 1% nonessential amino acids, penicillin (100 IU/mL), and streptomycin (100 μg/mL). 3. Serum-free medium: complete culture medium w/o FBS. 4. Maintenance conditions: humidified incubator under a 10% CO2/90% air atmosphere at 37
C (see Note 2). 5. Semi-permeable filters: 24 mm Transwell® (6-well plate), 3 μm pore size high-density (see Note 3).

18

Ba´rbara Graziela Postal et al.

2.2 Transepithelial Electrical Resistance (TEER)

1. 20–30 mL sterile 70% ethanol at room temperature. 2. 20–30 mL sterile 1 PBS w/o calcium and magnesium at room temperature. 3. 20–30 mL sterile serum-free medium at room temperature. 4. Volt-Ohm Meter (see Note 4).

2.3 Paracellular Permeability to Macromolecules

1. Sterile 96 wells black with clear flat bottom plate. 2. Sterile conical tubes (1.5 mL). 3. Sterile 200 μL and 1,000 μL pipette tips. 4. 40 mg/mL stock solution of FITC-labeled dextran 4 kDa (FD4) in serum-free medium, stored at 4
C (see Note 5). 5. Microplate fluorometer.

3

Methods All procedures should be carried out at room temperature and at a biosafety cabinet (BSC). All mediums should be at 37
C before use.

3.1

Cell Culture

1. Add 2.5 mL of complete medium in the basal compartment of each filter of a 6-well Transwell® plate (see Note 6) and seed Caco-2/TC7 cells on semi-permeable filters (apical compartment) at a density of 0.053  106 cells/cm2 (0.25  106 cells per filter of a 6-well Transwell® plate) in 1.5 mL of complete medium. Maintain the cells in an incubator with 10% CO2 at 37
C. 2. Two days after seeding, change the complete medium in both compartments (2 mL in basal compartment and 1 mL in apical compartment) and daily thereafter for 6 days. 3. From confluence (7 days after seeding in the conditions mentioned above), cells are switched to asymmetric conditions, i.e., with complete medium (2 mL) in the basal compartment and serum-free medium (1 mL) in the apical compartment (see Note 7). Both culture media are renewed daily for 14 days to reach an optimal enterocyte-like cell differentiation (see Note 8) [22, 23, 25, 27, 28]. 4. The day before measurement of paracellular permeability, renew the culture medium by adding 2.5 mL of complete medium in basal compartment and 1.5 mL of serum-free medium in apical compartment (see Note 9).

3.2 Transepithelial Electrical Resistance (TEER)

Transepithelial electrical resistance (TEER) can be used to follow the formation of the cell monolayer (TEER value increases from confluence) and to evaluate paracellular permeability modulations impacting the pore pathway (see Note 10).

Epithelial Paracellular Permeability in Caco-2/TC7 Cells

19

1. 24 h prior to the experiment: (a) Sterilize the electrode by immersing it for 10 min in a sterile 70% ethanol solution and leave the electrode overnight immersed in sterile PBS on the biosafety cabinet. Prepare 20–30 mL of sterile serum-free medium kept at room temperature overnight. (b) Prepare a plate with 2 filters (with the same characteristics that those used for the experiment) without cells and fill them with 2.5 mL of complete medium in the basal compartment and 1.5 mL of serum-free medium in the apical compartment (¼ null filters). Put them into the incubator (10% CO2 at 37
C) (see Note 11). (c) Change the medium of the filters containing Caco-2/ TC7 cells (2.5 mL complete medium in the basal compartment and 1.5 mL of serum-free medium in the apical compartment). 2. If the experiment is scheduled over several days and therefore the culture medium must be renewed daily, always measure the TEER before changing the medium (see Note 12). 3. Ten minutes prior to the measurements, equilibrate the electrode in the sterile serum-free medium at room temperature. 4. According to the instructions of the material used, perform a test of the Volt-Ohm Meter. 5. Take out the first plate and the plate with 2 null filters from the incubator. Leave these plates at room temperature on the biosafety cabinet for 7 min (use the timer) to stabilize the temperature of the culture medium (see Note 13). 6. Measure the resistance of the null filters, then the filters of the first plate of cells by placing the electrode successively in all the notches of each filter and put the plate back to the incubator. 7. Take out the second plate from the incubator and repeat the steps 5 and 6. The temperature equilibration of the second plate can be done during the measurement of the first plate, etc. 8. Calculate the TEER and express the results in ohm.cm2 as follows: (a) The resistance value of notches of each filter must be averaged, as well as the resistance values of null filters. (b) Subtract the average value of the null filters from the average value of each filter in the experiment and multiply by the membrane area of the filter (4.67 cm2 for a transwell® of a 6-well plate): (average Resistancefilter  average Resistancenull filter)  4.67 (see Note 14).

20

Ba´rbara Graziela Postal et al.

3.3 Paracellular Permeability to Macromolecules

Paracellular permeability across Caco-2/TC7 cell monolayer can be evaluated by measuring the accumulation during 4 h of FITC4 kDa dextran (FD4) in the basal compartment (see Note 10). 1. Turn off the biosafety cabinet light as soon as you use the FITC-labeled dextran. 2. Prepare sufficient solution (1.5 mL per well) for all filters of FITC-labeled dextran 4 kDa (FD4) solution at 1 mg/mL in serum-free medium. 3. Replace the apical medium by 1 mg/mL FD4 in serum-free medium (1.5 mL per filter) (see Note 15). Do not change the basal medium (see Note 16). 4. Incubate in 10% CO2 at 37
C for 2 h. Be careful not to shake the plate when removing it or putting it back in the incubator. 5. Using a single-channel 200 μL pipette, set to 115 μL, gently insert the cone into a notch, make a go back and forth to homogenize the basal medium, take 115 μL and put into the conical tube (1.5 mL) corresponding to the well, with the same pipette tip put on in another notch of the same filter, make an another go back and forth, retake 115 μL and put it into the same conical tube. Be careful not to move the filter when collecting the basal medium samples. In the end, every conical tube (1.5 mL) contains 2  115 μL ¼ 230 μL. Go to the next filter and repeat the procedure. These samples are the 2-h measurements. Protect the tubes from light at 4
C. 6. When the basal medium of all filters is collected, with a 1000 μL pipette, add 230 μL of complete medium warmed to 37
C in one notch per filter. Do not go back and forth. Put back into incubator in 10% CO2 at 37
C for 2 h. 7. Repeat the step 5. These samples are the 4-h measurements (see Note 17). 8. Prepare the FD4 standard concentrations (protect it from the light): Dilute in complete medium the remaining 1 mg/mL FD4 solution prepared for the experiment to obtain the following concentrations: (10 μg/mL; 7.5 μg/mL; 5 μg/mL; 2.5 μg/mL; 1 μg/mL; 0.75 μg/mL; 0.5 μg/mL; 0.25 μg/ mL; 0.1 μg/mL; 0.5 μg/mL; 0.01 μg/mL). The standard concentration of 10 μg/mL corresponds to 1% of the initial concentration of FD4 in the apical compartment. 9. In sterile assay plate, 96 wells black with clear flat bottom: (a) Add 120 μL from the standard range (from 10 to 0.01 μg/mL) in duplicate. (b) Add 120 μL from the complete medium alone, as blank to subtract the basal medium background fluorescence emission.

Epithelial Paracellular Permeability in Caco-2/TC7 Cells

21

(c) Add 120 μL from each sample collected at time 2 h and 4 h. 10. Determine the fluorescence values with a microplate fluorometer. Read the fluorescence at the wavelength 485 nm excitation and 535 nm emission. Set the gain on the well corresponding to the 5 μg/mL standard range concentration (see Note 18). 11. Use the regression line to determine at each time (2 h and 4 h) the FD4 concentration in the basal compartment. Results can be expressed as the percentage of FD4 initially added in the apical compartment (see Note 19).

4

Notes 1. Caco-2/TC7 cell line is available under request to Ve´ronique Carrie`re or Sophie Thenet who are in charge of this cell line since the retirement of Monique Rousset. All uses of Caco-2/ TC7 by a non-academic entity require a license from INSERMtransfert. 2. DMEM used for Caco-2/TC7 cells containing 3.7 g/L sodium bicarbonate, the cells are cultured in a 10% CO2 atmosphere. However, Caco-2/TC7 cells can also grow and differentiate in a 5% CO2 incubator but it is worth noting that such condition impacts metabolic activities of the cells in particular glucose metabolism (personal observations). 3. Various microplate sizes (6, 12, and 24 wells) and pore sizes membranes (i.e., 3 μm, 3 μm high-density, 1 μm, 0.4 μm) can be used. The nature of filter membrane can affect the absolute values of measured paracellular permeability [26, 29]. 4. There are different TEER measurement methods, the Ohm’s law method, and the impedance spectroscopy. The widely used and commercially available TEER measurement system is based in the Ohm’s law method, known as an Epithelial Volt-Ohm Meter (EVOM). The TEER readings with EVOM can be realized in manual and/or automatic system. 5. The mechanisms that regulate paracellular permeability through the different pathways are complex and it may be important to explore the variations of paracellular permeability to molecules of different sizes such 10 kDa-, 40 kDa-, 70 kDadextran or sulfonic acid (0.4 kDa) [30–32]. Although the size limit of molecules that can cross an epithelium through the leak pathway is still debated, larger molecules such as 40 kDa-dextran and over are considered by some authors to be excluded from the paracellular pathways and are commonly used to evaluate transcellular pathway. It is likely that a mixture of

22

Ba´rbara Graziela Postal et al.

paracellular and transcellular pathways is evaluated using large tracers, depending on the experimental conditions (see chapter “Measure of paracellular permeability in mouse intestine using Ussing chambers” by Aguanno et al. in this book). 6. Cell seeding is more homogenous when the plates are filled with the complete medium in the basal compartment and placed in the CO2 incubator 10 min before plating cells. 7. Asymmetric culture conditions help to reproduce the physiological environment of intestinal epithelial cells. In particular, serum-free culture medium in the apical compartment mimics the intestinal lumen, which is devoid of serum. 8. From confluence, Caco-2/TC7 cells start to differentiate and polarize. They form a cell monolayer with an efficient selective barrier. The presence of phenol red in the culture medium helps to visually estimate the integrity of the monolayer. Indeed the secretion of lactic acid at the basal pole of polarized cells decreases the pH of the basal medium and turns it orange while the apical medium remains red. 9. Increasing the volume of culture medium ensures a good immersion of the electrodes for TEER measurement. If only paracellular permeability to macromolecules is assayed, the volume of culture medium can be kept at 2 mL (basal) and 1 mL (apical). 10. EGTA can be used as a positive control of increased paracellular permeability affecting both pore and leak pathways [23, 25]. EGTA is a calcium chelator known to provoke the disruption of junctional protein complexes and to prevent the return of junction proteins to the membrane, resulting in the opening of the intercellular junctions [33]. Addition of cytokines in the basal compartment can be used as a positive control for increased paracellular permeability to macromolecules (leak pathway) and, hence, FD4 passage [22, 34]. 11. The null filters (without cells) are used to measure the intrinsic resistance of the filter. 12. It is important to note that TEER measurements are sensitive to changes in the composition of the culture medium. The renewal of the medium before the measurement of TEER can affect the results. 13. TEER measurements have been reported to be temperature dependent. The temperature should be equilibrated to room temperature before performing resistance measurements to avoid any temperature fluctuation-induced TEER changes. However, the time out of incubator could be detrimental to cell physiology and function, thus 7 min waiting time is the time which seems a good compromise for stabilization of the

Epithelial Paracellular Permeability in Caco-2/TC7 Cells

23

TEER values without removing the cells from the incubator for too long [26]. 14. Untreated Caco-2/TC7 cells cultured for 21 days on 3 μm high-density transwell® of 6-well plate display a TEER value between 600 and 800 Ω.cm2 [22, 23, 25]. 15. Since fluorescence variations are very sensitive, it is better to use the same preparation of 1 mg/mL FD4 solution for all wells. When different treatments are added in the apical compartments, split the FD4 solution in separate tubes and add the molecules/drugs at the appropriate final concentrations before changing the apical medium of the cells. 16. Serum in the complete medium brings growth factors that can trigger unwanted transient modification of paracellular permeability. The non-renewal of the basal medium on the day of the experiment increases the strength and the reproducibility of the results. If an experimental treatment must be supplied in the basal compartment the day of the experiment, it is better to add concentrate solution of the drug tested without renewing the basal medium. 17. Evaluation of FD4 accumulation at two different time points (2 h and 4 h) helps to control the integrity of cell monolayer throughout the experiment. A 1.5-fold increase in FD4 accumulation in untreated Caco-2/TC7 cells is generally observed between 2 h and 4 h. 18. Usually the standard concentration 5 μg/mL is used for setting the gain, since FD4 passage rarely exceeds 0.5% of the initial amount. In this case do not use the 10 μg/L and 7.5 μg/mL standard range concentrations to make the regression line. Otherwise a gain can be calculated taking into account all the wells. 19. For samples collected at time-point 2 h, the quantity (μg) of FD4 present in the basal compartment is calculated as follows: (FD4 concentration  2.5 mL). For samples collected at timepoint 4 h, take into account the μg of FD4 removed in the 0.23 mL collected at time 2 h. The final quantity (μg) in samples at time 4 h is calculated for each transwell® as follows: (FD4 concentration in samples at 4 h  2.5 mL) + (FD4 concentration of the same filter at 2 h  0.23 mL). To express the results as percentage of initial amount of FD4 placed in apical compartment, use the following formulation: (FD4 quantity in μg calculated above  100)/1500. A typical value of FD4 passage in the basal compartment obtained at time 4 h in untreated Caco-2/TC7 cells cultured for 21 days on 3μmHD transwell® is 0.03%.

24

Ba´rbara Graziela Postal et al.

Acknowledgments This work was supported by: the Brazilian government’s Science Without Borders Program, the Association Franc¸ois Aupetit (AFA), Institut National de la Sante´ et de la Recherche Me´dicale, Sorbonne Universite´, Ecole Pratique des Hautes Etudes. B.G.P. received a doctoral fellowship (CNPq 207303/2014-2). D.A. received a fellowship from CORDDIM Ile de France. References 1. Claude P, Goodenough DA (1973) Fracture faces of zonulae occludentes from “tight” and “leaky” epithelia. J Cell Biol 58(2):390–400. https://doi.org/10.1083/jcb.58.2.390 2. Fro¨mter E, Diamond J (1972) Route of passive ion permeation in epithelia. Nat New Biol 235 (53):9–13. https://doi.org/10.1038/ newbio235009a0 3. Machen TE, Erlij D, Wooding FB (1972) Permeable junctional complexes. The movement of lanthanum across rabbit gallbladder and intestine. J Cell Biol 54(2):302–312. https:// doi.org/10.1083/jcb.54.2.302 4. Groschwitz KR, Hogan SP (2009) Intestinal barrier function: molecular regulation and disease pathogenesis. J Allergy Clin Immunol 124 (1):3–20; quiz 21-22. https://doi.org/10. 1016/j.jaci.2009.05.038 5. Lee JY, Wasinger VC, Yau YY, Chuang E, Yajnik V, Leong RW (2018) Molecular pathophysiology of epithelial barrier dysfunction in inflammatory bowel diseases. Proteomes 6 (2):17. https://doi.org/10.3390/ proteomes6020017 6. Suzuki T (2013) Regulation of intestinal epithelial permeability by tight junctions. Cell Mol Life Sci 70(4):631–659. https://doi. org/10.1007/s00018-012-1070-x 7. Le Gall M, Thenet S, Aguanno D, Jarry AC, Genser L, Ribeiro-Parenti L, Joly F, Ledoux S, Bado A, Le Beyec J (2019) Intestinal plasticity in response to nutrition and gastrointestinal surgery. Nutr Rev 77(3):129–143. https:// doi.org/10.1093/nutrit/nuy064 8. Turner JR (2009) Intestinal mucosal barrier function in health and disease. Nat Rev Immunol 9(11):799–809. https://doi.org/10. 1038/nri2653 9. Shen L, Weber CR, Raleigh DR, Yu D, Turner JR (2011) Tight junction pore and leak pathways: a dynamic duo. Annu Rev Physiol 73:283–309. https://doi.org/10.1146/ annurev-physiol-012110-142150

10. Fogh J, Fogh JM, Orfeo T (1977) One hundred and twenty-seven cultured human tumor cell lines producing tumors in nude mice. J Natl Cancer Inst 59(1):221–226. https://doi. org/10.1093/jnci/59.1.221 11. Pinto M, Robine-Leon S, Appay M-D, Kedinger M, Haffen K, Fogh J, Zweibaum A (1983) Enterocyte-like differentiation and polarization of the human colon carcinoma cell line Caco-2 in culture. Biol Cell 47:323–330 12. Hauri HP, Sterchi EE, Bienz D, Fransen JA, Marxer A (1985) Expression and intracellular transport of microvillus membrane hydrolases in human intestinal epithelial cells. J Cell Biol 101(3):838–851. https://doi.org/10.1083/ jcb.101.3.838 13. Zweibaum A, Triadou N, Kedinger M, Augeron C, Robine-Le´on S, Pinto M, Rousset M, Haffen K (1983) Sucraseisomaltase: a marker of foetal and malignant epithelial cells of the human colon. Int J Cancer 32(4):407–412. https://doi.org/10.1002/ ijc.2910320403 14. Fois CAM, Le TYL, Schindeler A, Naficy S, McClure DD, Read MN, Valtchev P, Khademhosseini A, Dehghani F (2019) Models of the gut for analyzing the impact of food and drugs. Adv Healthc Mater 8 (21):1900968. https://doi.org/10.1002/ adhm.201900968 15. Zweibaum A, Laburthe M, Grasset E, Louvard D (1991) Use of cultured cell lines in studies of intestinal cell differentiation and function. In: Intestinal absorption and secretion: handbook of physiology, section 6, the gastrointestinal system. American Physiological Society, Bethesda, pp 223–255 16. Herrera-Ruiz D, Wang Q, Cook TJ, Knipp GT, Gudmundsson OS, Smith RL, Faria TN (2001) Spatial expression patterns of peptide transporters in the human and rat gastrointestinal tracts, Caco-2 in vitro cell culture model,

Epithelial Paracellular Permeability in Caco-2/TC7 Cells and multiple human tissues. AAPS PharmSci 3 (1):100. https://doi.org/10.1208/ps030109 17. Jung D, Fried M, Kullak-Ublick GA (2002) Human apical sodium-dependent bile salt transporter gene (SLC10A2) is regulated by the peroxisome proliferator-activated receptor α. J Biol Chem 277(34):30559–30566. https://doi.org/10.1074/jbc.M203511200 18. Artursson P, Magnusson C (1990) Epithelial transport of drugs in cell culture. II: effect of extracellular calcium concentration on the paracellular transport of drugs of different lipophilicities across monolayers of intestinal epithelial (Caco-2) cells. J Pharm Sci 79 (7):595–600. https://doi.org/10.1002/jps. 2600790710 19. Hidalgo IJ, Raub TJ, Borchardt RT (1989) Characterization of the human colon carcinoma cell line (Caco-2) as a model system for intestinal epithelial permeability. Gastroenterology 96(3):736–749 20. Chantret I, Rodolosse A, Barbat A, Dussaulx E, Brot-Laroche E, Zweibaum A, Rousset M (1994) Differential expression of sucraseisomaltase in clones isolated from early and late passages of the cell line Caco-2: evidence for glucose-dependent negative regulation. J Cell Sci 107(Pt 1):213–225 21. Zucco F, Batto A-F, Bises G, Chambaz J, Chiusolo A, Consalvo R, Cross H, Dal Negro G, de Angelis I, Fabre G, Guillou F, Hoffman S, Laplanche L, Morel E, Pinc¸onRaymond M, Prieto P, Turco L, Ranaldi G, Rousset M, Sambuy Y, Scarino ML, Torreilles F, Stammati A (2005) An interlaboratory study to evaluate the effects of medium composition on the differentiation and barrier function of Caco-2 cell lines. Altern Lab Anim 33(6):603–618. https://doi.org/ 10.1177/026119290503300618 22. Aguanno D, Coquant G, Postal BG, Osinski C, Wieckowski M, Stockholm D, Grill JP, Carrie`re V, Seksik P, Thenet S (2020) The intestinal quorum sensing 3-oxo-C12:2 acyl homoserine lactone limits cytokine-induced tight junction disruption. Tissue Barriers 8 (4):1832877. https://doi.org/10.1080/ 21688370.2020.1832877 23. Ghezzal S, Postal BG, Quevrain E, Brot L, Seksik P, Leturque A, Thenet S, Carriere V (2020) Palmitic acid damages gut epithelium integrity and initiates inflammatory cytokine production. Biochim Biophys Acta Mol Cell Biol Lipids 1865(2):158530. https://doi. org/10.1016/j.bbalip.2019.158530 24. Petit CS, Barreau F, Besnier L, Gandille P, Riveau B, Chateau D, Roy M, Berrebi D, Svrcek M, Cardot P, Rousset M, Clair C,

25

Thenet S (2012) Requirement of cellular prion protein for intestinal barrier function and mislocalization in patients with inflammatory bowel disease. Gastroenterology 143 (1):122–132.e15. https://doi.org/10.1053/ j.gastro.2012.03.029 25. Postal BG, Ghezzal S, Aguanno D, Andre´ S, Garbin K, Genser L, Brot-Laroche E, Poitou C, Soula H, Leturque A, Cle´ment K, Carrie`re V (2020) AhR activation defends gut barrier integrity against damage occurring in obesity. Mol Metab 39:101007. https://doi. org/10.1016/j.molmet.2020.101007 26. Srinivasan B, Kolli AR, Esch MB, Abaci HE, Shuler ML, Hickman JJ (2015) TEER measurement techniques for in vitro barrier model systems. J Lab Autom 20(2):107–126. https://doi.org/10.1177/ 2211068214561025 27. Beaslas O, Cueille C, Delers F, Chateau D, Chambaz J, Rousset M, Carriere V (2009) Sensing of dietary lipids by enterocytes: a new role for SR-BI/CLA-1. PLoS One 4(1):e4278 28. Morel E, Ghezzal S, Lucchi G, Truntzer C, Pais de Barros JP, Simon-Plas F, Demignot S, Mineo C, Shaul PW, Leturque A, Rousset M, Carriere V (2018) Cholesterol trafficking and raft-like membrane domain composition mediate scavenger receptor class B type 1-dependent lipid sensing in intestinal epithelial cells. Biochim Biophys Acta 1863 (2):199–211. https://doi.org/10.1016/j. bbalip.2017.11.009 29. Srinivasan B, Kolli AR (2019) Transepithelial/ transendothelial electrical resistance (TEER) to measure the integrity of blood-brain barrier. In: Barichello T (ed) Blood-brain barrier. Springer New York, New York, NY, pp 99–114 30. Al-Sadi R, Khatib K, Guo S, Ye D, Youssef M, Ma T (2011) Occludin regulates macromolecule flux across the intestinal epithelial tight junction barrier. Am J Physiol Gastrointest Liver Physiol 300(6):G1054–G1064. https:// doi.org/10.1152/ajpgi.00055.2011 31. Genser L, Aguanno D, Soula HA, Dong L, Trystram L, Assmann K, Salem JE, Vaillant JC, Oppert JM, Laugerette F, Michalski MC, Wind P, Rousset M, Brot-Laroche E, Leturque A, Cle´ment K, Thenet S, Poitou C (2018) Increased jejunal permeability in human obesity is revealed by a lipid challenge and is linked to inflammation and type 2 diabetes. J Pathol 246(2):217–230. https://doi. org/10.1002/path.5134 32. Ikenouchi J, Furuse M, Furuse K, Sasaki H, Tsukita S, Tsukita S (2005) Tricellulin constitutes a novel barrier at tricellular contacts of

26

Ba´rbara Graziela Postal et al.

epithelial cells. J Cell Biol 171(6):939–945. https://doi.org/10.1083/jcb.200510043 33. Tria S, Jimison LH, Hama A, Bongo M, Owens RM (2013) Sensing of EGTA mediated barrier tissue disruption with an organic transistor. Biosensors 3(1):44–57. https://doi.org/10. 3390/bios3010044

34. Wang F, Graham WV, Wang Y, Witkowski ED, Schwarz BT, Turner JR (2005) Interferongamma and tumor necrosis factor-alpha synergize to induce intestinal epithelial barrier dysfunction by up-regulating myosin light chain kinase expression. Am J Pathol 166 (2):409–419. https://doi.org/10.1016/ s0002-9440(10)62264-x

Methods in Molecular Biology (2021) 2367: 27–35 DOI 10.1007/7651_2021_346 © Springer Science+Business Media, LLC 2021 Published online: 05 March 2021

Evaluation of Barrier Functions in Human iPSC-Derived Intestinal Epithelium Shigeru Yamada and Yasunari Kanda Abstract The small intestine plays roles in the absorption and metabolism of orally administered drugs and chemicals. Tight junctions between intestinal epithelial cells, which form a tight barrier preventing the invasion of pathogens and toxins, are essential components of the intestinal defense system. These intestinal functions have generally been evaluated using established cell lines or primary cells in two-dimensional culture. However, these culture systems have not shown the complexity of the three-dimensional structure and diversity of cell types comprising the intestinal epithelial tissue. Here, we report the generation of intestinal organoids using human induced pluripotent stem cells subjected to sequential treatment with different cytokines and compounds. We further describe the tool for evaluating intestinal barrier functions using organoids as a physiologically relevant human platform. Key words Human iPS cells, Intestine, Organoid, Tight junction, Barrier

1

Introduction In humans, orally ingested drugs and chemicals are primarily absorbed in the intestine. Intestinal epithelial cells form a tight barrier between the gut and the intestinal lumen, thereby preventing the invasion of pathogens and toxins into the human body, while retaining the capacity to absorb compounds processed by drug-metabolizing enzymes [1, 2]. Thus, appropriate epithelial barrier maturation is essential for intestinal defense and homeostasis. As a model of the intestinal epithelium, human colon cancer Caco-2 cells have been widely used for assessing intestinal barrier functions with respect to different compounds. Previous studies have indicated certain changes in epithelial barrier properties, including an increase in transepithelial electrical resistance (TEER: an indicator of epithelial barrier integrity) and a reduction in paracellular permeability [3]. In addition, Caco-2 barrier integrity has been reported to be disrupted by mycotoxins, including aflatoxin, ochratoxin A, patulin, and zearalenone [4]. However, concerns have been raised regarding the use of Caco-2 cells as a model of the human intestine. For example, the TEER values of Caco-2 cells

27

28

Shigeru Yamada and Yasunari Kanda

have been reported to be higher than those observed in normal human intestinal cells [5], suggesting the possibility of an abnormal tightness between the cells. Differences between Caco-2 cells and the human intestine have also been observed with respect to transporter levels and permeability [6]. Accordingly, given these inconsistencies, it would be desirable to establish alternative novel in vitro models for the evaluation of compounds. Human induced pluripotent stem cells (iPSCs) have the potential to differentiate into almost all cell types that constitute the human body [7] and are viewed as providing a valuable tool in the assessment of diverse compounds of interest [8, 9]. In this regard, numerous recent studies have examined three-dimensional (3D) organoid models based on human iPSCs subjected to treatment with cytokines and chemicals [10–17]. Matrigel overlay protocol, for example, can be used to mimic human intestinal development at the fetal stage [10, 11, 14–17]. These intestinal organoids have morphological characteristics of intestinal tissues, such as a polarized epithelium, which forms villus-crypt structures, and contain multiple epithelial cell types, including nutrientabsorbing enterocytes, goblet cells, paneth cells, and intestinal stem cells [10, 11, 14–17]. The intestinal models based on iPSCs are considered to have the potential to overcome disadvantages associated with the use of Caco-2 cells. In this study, we describe a protocol for generation of intestinal organoids from human iPSCs, and their use in the evaluation of intestinal barrier functions.

2

Materials 1. Human iPSC line 253G1 (Riken BRC Cell Bank, Ibaraki, Japan). 2. TeSR-E8 medium (Stemcell Technologies, Vancouver, BC, Canada). 3. Human embryonic stem cell-qualified (BD Biosciences, San Jose, CA, USA). 4. Accumax (Innovative CA, USA).

Cell

Technologies,

Matrigel

San

5. Y-27632 (Wako, Tokyo, Japan). 6. RPMI1640 medium (Nacalai Tesque, Kyoto, Japan). 7. CHIR99021 (Selleck Chemicals, Houston, TX, USA). 8. Activin A (R&D Systems, Minneapolis, MN, USA). 9. B27 (Thermo Fisher Scientific, Waltham, MA, USA). 10. Fibroblast growth factor 4 (FGF4; R&D Systems). 11. Advanced DMEM:F12 (Thermo Fisher Scientific).

Diego,

Evaluation of iPSC-Derived Intestinal Barrier Functions

29

12. N2 (Thermo Fisher Scientific). 13. Epidermal growth factor (EGF; R&D Systems). 14. Noggin (R&D Systems). 15. R-spondin-1 (R&D Systems). 16. Cell recovery solution (BD Biosciences). 17. TrypLE Select (Thermo Fisher Scientific). 18. Hepatocyte growth factor (HGF; R&D Systems). 19. Wnt3a (R&D Systems). 20. SB202190 (Sigma-Aldrich, St. Louis, MO, USA). 21. A83-01 (Tocris, Bristol, UK). 22. Type I collagen (Nippi, Tokyo, Japan). 23. 24-well transwell plate (Merck, Poole, UK). 24. 40 ,6-diamidino-2-phenylindole (DAPI, Nacalai Tesque). 25. SlowFade (Thermo Fisher Scientific). 26. BIOREVO BZ-9000 fluorescence microscope (Keyence, Osaka, Japan). 27. Confocal laser scanning microscope (Nikon A1; Nikon, Tokyo, Japan). 28. Millicell ERS-2 MA, USA).

volt-ohm

meter

(Millipore,

Bedford,

29. Fluorescein isothiocyanate (FITC)-dextran (Sigma-Aldrich). 30. Fluoroskan Ascent FL microplate reader (Thermo Fisher Scientific). 31. TRIzol (Thermo Fisher Scientific). 32. QuantiTect SYBR Green RT-PCR Kit (Qiagen, Valencia, CA, USA). 33. QuantStudio 7 Flex Real-Time PCR System (Thermo Fisher Scientific). 34. Antibodies for immunocytochemistry (see Table 1). 35. Primers for real-time PCR (see Table 2).

3

Methods

3.1 Differentiation of Human iPSCs into Intestinal Organoids

1. Prepare iPSCs at around 90% confluence (see Note 1). 2. The iPSCs were differentiated into definitive endoderm (DE) by incubation in RPMI1640 medium containing B27, CHIR99021 (2 μM), and activin A (100 ng/mL) for 24 h, followed by incubation in RPMI1640 containing B27 and activin A for 48 h.

30

Shigeru Yamada and Yasunari Kanda

Table 1 Antibodies for immunocytochemistry Name

Dilution

Supplier

SOX17

1:500

R&D Systems

FOXA2

1:1000

Cell Signaling Technology

CDX2

1:500

Thermo Fisher Scientific

ECAD

1:50

Cell Signaling Technology

KLF5

1:100

Proteintech

ZO-1

1:400

Cell Signaling Technology

Table 2 Primers for real-time PCR Gene

Forward primer

Reverse primer

ECAD

ATTTTTCCCTCGACACCCGAT

TCCCAGGCGTAGACCAAGA

KLF5

CATCCACTACTGCGATTACCC

CCCAGGTACACTTGTATGGC

CDX2

GGAACCTGTGCGAGTGGAT

TCGATATTTGTCTTTCGTCCTG

LGR5

CACCTCCTACCTAGACCTCAGT

CGCAAGACGTAACTCCTCCAG

VIL1

CGGAAAGCACCCGTATGGAG

CGTCCACCACGCCTACATAG

MUC2

GAGGGCAGAACCCGAAACC

GGCGAAGTTGTAGTCGCAGAG

LYZ

CCCTGGTCAGCCTAGCACTC

CCTTGCCCTGGACCGTAACA

CYP3A4

AAGTCGCCTCGAAGATACACA

AAGGAGAGAACACTGCTCGTG

ZO-1

CAACATACAGTGACGCTTCACA

CACTATTGACGTTTCCCCACTC

GAPDH

GTCTCCTCTGACTTCAACAGCG

ACCACCCTGTTGCTGTAGCCAA

3. DE cells were further differentiated into mid/hindgut by incubation in RPMI1640 containing B27, CHIR99021, activin A, and FGF4 (500 ng/mL) for up to 4 days (see Note 2). 4. Free-floating spheroids were collected during the course of culture, transferred to Matrigel, and provided with intestinal growth medium [Advanced DMEM:F12 containing B27, N2, EGF (100 ng/mL), noggin (100 ng/mL), and R-spondin-1 (500 ng/mL)] (see Note 3). 5. The medium was changed at 2–3 days intervals over a period of 14 days [17] (Fig. 1). 3.2 Monolayer Culture from Intestinal Organoids

1. The organoids embedded in Matrigel were washed with phosphate-buffered saline (PBS) and treated with cell recovery solution for 40 min at 4  C (see Note 4).

Evaluation of iPSC-Derived Intestinal Barrier Functions

31

A Day0

Day1

Act A CHIR

Day3

Day7

Act A CHIR FGF4

Act A

Day21

EGF RSPO1 NOG

B ECAD

KLF5

DAPI

DIC

Bar=20mm

Fig. 1 Differentiation into human iPSC-derived intestinal organoids. (a) A schematic representation of the procedures used for intestinal organoid differentiation from human iPSCs. CHIR CHIR99021, Act A activin A, RSPO1 R-spondin-1, NOG noggin. (b) Intestinal organoids (Day 23) after the differentiation. The localization of marker proteins was analyzed by immunocytochemistry. Nuclei were counterstained with DAPI. DIC differential interference contrast. Bar ¼ 20 μm

2. Collect the recovered organoids by centrifugation at 200  g for 3 min. 3. Incubate the organoids in TrypLE Select containing Y-27632 for 15 min at 37  C, and dissociate into single cells by pipetting (see Note 5). 4. After centrifugation at 200  g for 5 min, the pelleted cells were resuspended in organoid growth medium [Advanced DMEM:F12 containing B27 (minus vitamin A), N2, EGF, noggin, R-spondin-1, HGF (50 ng/mL), Wnt3a (50 ng/ mL), SB202190 (10 μM), A83-01 (500 nM), 1% bovine serum albumin, and Y-27632]. 5. Seed in a Type I collagen-coated 24-well transwell plate (see Note 6). 3.3 Measurement of TEER

1. Monolayer cells of transwell cultures were washed twice in Advanced DMEM:F12, and allowed to equilibrate for 15 min at room temperature (see Note 7). 2. Cell monolayer TEER was measured in Advanced DMEM: F12 at room temperature using an Endohm tissue resistance chamber connected to a Millicell ERS-2 volt-ohm meter. 3. The resistance of Advanced DMEM:F12 alone was considered as background resistance and subtracted from the measured TEER values [17] (Figs. 2 and 3).

Shigeru Yamada and Yasunari Kanda

dispersed cells

transwell insert

collagen-coated polyester membrane

multiwell plate

TEER Permeability

Tight junction Fig. 2 Schematic diagram of transepithelial barrier function assays

)

150

100

TEER (Ω

32

50

0 0

2

4

6

8

10

Time course (day) Fig. 3 Time course studies of TEER value. After seeding and culturing in transwell chambers for 2, 5, and 10 days, each TEER value of intestinal epithelium was measured

Evaluation of iPSC-Derived Intestinal Barrier Functions

3.4 Permeability Assay

33

1. The membrane-impermeable tracer FITC-dextran, with a molecular mass of 40 kDa, was dissolved in Hank’s balanced salt solution (pH 7.4) at a final concentration of 100 μg/mL. 2. The tracer was added to the apical monolayer cells in transwells for 2 h. Transwells without monolayer cells were used as controls. 3. The fluorescence intensity of permeabilized dextran in the basal chamber was measured using a Fluoroskan Ascent FL microplate reader with excitation and emission at 488 nm and 515 nm, respectively [17] (Fig. 2).

3.5 Assessment of Tight Junction Formation (ZO-1 Staining)

1. Cells on a transwell insert membrane were fixed in 4% paraformaldehyde for 20 min at room temperature. 2. Permeabilize the cells with 0.5% Triton X-100 in PBS (pH 7.4). 3. Block the cells with 10% FBS and incubate overnight with primary antibodies against ZO-1 (1:400) at 4  C. 4. Incubate the cells with an appropriate fluorescent secondary antibody at 4  C. 5. Cell nuclei were counterstained with DAPI. 6. Cut off a transwell insert membrane with a utility knife (see Note 8). 7. Cells were mounted in SlowFade and examined under a confocal laser scanning microscope [17] (Fig. 2).

4

Notes 1. The density of iPSCs prior to induction is critical for the subsequent differentiation processes. 2. After approximately 48 h from the start of mid/hindgut differentiation, spheroids became visible. 3. The supernatant containing free-floating spheroids is conjugated with Matrigel (25 μL supernatant + 50 μL Matrigel) and then added dropwise to the middle of wells. 4. After an initial incubation for 20 min, shake the culture vessel gently for better cell recovery. 5. This step is critical for monolayer epithelial formation on transwell insert membranes. The organoids should be thoroughly dissociated into single cells. 6. Considering the difficulty to observe the cells on transwell insert membranes under the microscope, the remaining cells should be seeded in 96-well plates for checking. 7. Care should be exercised to avoid damaging the cell sheet when changing the medium. 8. Attention should be paid to both the upper and lower surfaces of the membrane.

34

Shigeru Yamada and Yasunari Kanda

Acknowledgements This work was supported by a Grant-in-Aid for Scientific Research from the Ministry of Education, Culture, Sports, Science, and Technology, Japan (#17H04107 and #17K19503 to Y. K., #17K00576 to S. Y.), the Research on Regulatory Harmonization and Evaluation of Pharmaceuticals, Medical Devices, Regenerative and Cellular Therapy Products, Gene Therapy Products, and Cosmetics from Japan Agency for Medical Research and Development, AMED (JP18mk0104117 to Y. K.), and a grant from the Smoking Research Foundation (Y. K.). References 1. Kelly JR, Kennedy PJ, Cryan JF, Dinan TG, Clarke G, Hyland NP (2015) Breaking down the barriers: the gut microbiome, intestinal permeability and stress-related psychiatric disorders. Front Cell Neurosci 9:392 2. Peters SA, Jones CR, Ungell AL, Hatley OJ (2016) Predicting drug extraction in the human gut wall: assessing contributions from drug metabolizing enzymes and transporter proteins using preclinical models. Clin Pharmacokinet 55(6):673–696 3. Valenzano MC, DiGuilio K, Mercado J, Teter M, To J, Ferraro B, Mixson B, Manley I, Baker V, Moore BA, Wertheimer J, Mullin JM (2015) Remodeling of tightjunctions and enhancement of barrier integrity of the CACO-2 intestinal epithelial cell layer by micronutrients. PLoS One 10(7):e0133926 4. Akbari P, Braber S, Varasteh S, Alizadeh A, Garssen J, Fink-Gremmels J (2017) The intestinal barrier as an emerging target in the toxicological assessment of mycotoxins. Arch Toxicol 91(3):1007–1029 5. van Breemen RB, Li Y (2005) Caco-2 cell permeability assays to measure drug absorption. Expert Opin Drug Metab Toxicol 1 (2):175–185 6. Sun D, Lennernas H, Welage LS, Barnett JL, Landowski CP, Foster D, Fleisher D, Lee KD, Amidon GL (2002) Comparison of human duodenum and Caco-2 gene expression profiles for 12,000 gene sequences tags and correlation with permeability of 26 drugs. Pharm Res 19(10):1400–1416 7. Li K, Kong Y, Zhang M, Xie F, Liu P, Xu S (2016) Differentiation of pluripotent stem cells for regenerative medicine. Biochem Biophys Res Commun 471(1):1–4 8. Kanda Y, Yamazaki D, Osada T, Yoshinaga T, Sawada K (2018) Development of

torsadogenic risk assessment using human induced pluripotent stem cell-derived cardiomyocytes: Japan iPS Cardiac Safety Assessment (JiCSA) update. J Pharmacol Sci 138 (4):233–239 9. Yamada S, Kubo Y, Yamazaki D, Sekino Y, Kanda Y (2017) Chlorpyrifos inhibits neural induction via Mfn1-mediated mitochondrial dysfunction in human induced pluripotent stem cells. Sci Rep 7:40925 10. Clinton J (2016) Directed differentiation of gastrointestinal epithelial organoids using ATCC CELLMATRIX basement membrane from multiple human ATCC iPSC lines. AP Notes 26:1–8 11. McCracken KW, Howell JC, Wells JM, Spence JR (2011) Generating human intestinal tissue from pluripotent stem cells in vitro. Nat Protoc 6(12):1920–1928 12. Negoro R, Takayama K, Nagamoto Y, Sakurai F, Tachibana M, Mizuguchi H (2016) Modeling of drug-mediated CYP3A4 induction by using human iPS cell-derived enterocyte-like cells. Biochem Biophys Res Commun 472(4):631–636 13. Ogaki S, Morooka M, Otera K, Kume S (2015) A cost-effective system for differentiation of intestinal epithelium from human induced pluripotent stem cells. Sci Rep 5:17297 14. Onozato D, Yamashita M, Nakanishi A, Akagawa T, Kida Y, Ogawa I, Hashita T, Iwao T, Matsunaga T (2018) Generation of intestinal organoids suitable for pharmacokinetic studies from human induced pluripotent stem cells. Drug Metab Dispos 46 (11):1572–1580 15. Spence JR, Mayhew CN, Rankin SA, Kuhar MF, Vallance JE, Tolle K, Hoskins EE, Kalinichenko VV, Wells SI, Zorn AM, Shroyer NF, Wells JM (2011) Directed differentiation of

Evaluation of iPSC-Derived Intestinal Barrier Functions human pluripotent stem cells into intestinal tissue in vitro. Nature 470(7332):105–109 16. Uchida H, Machida M, Miura T, Kawasaki T, Okazaki T, Sasaki K, Sakamoto S, Ohuchi N, Kasahara M, Umezawa A, Akutsu H (2017) A xenogeneic-free system generating functional

35

human gut organoids from pluripotent stem cells. JCI Insight 2(1):e86492 17. Yamada S, Kanda Y (2019) Retinoic acid promotes barrier functions in human iPSC-derived intestinal epithelial monolayers. J Pharmacol Sci 140(4):337–344

Methods in Molecular Biology (2021) 2367: 37–46 DOI 10.1007/7651_2020_313 © Springer Science+Business Media New York 2020 Published online: 14 August 2020

Selective Regional Isolation of Brain Microvessels Fernanda Medina-Flores, Gabriela Hurtado-Alvarado, and Beatriz Go´mez-Gonza´lez Abstract The study of the regionalized function of the blood-brain barrier at the level of brain endothelial cells and pericytes is essential to understand the biological properties and molecular mechanisms regulating this biological barrier. The isolation of blood vessels from specific brain regions will allow to understand regional differences in susceptibility to pathological phenomena such as ischemia, traumatic brain injury, and neurodegenerative diseases, such as Alzheimer disease. Here, we propose an efficient and fast method to isolate brain endothelial cells and pericytes from a specific cerebral region. The isolated brain endothelial cells and pericytes are viable to perform conventional molecular and histological techniques such as Western blots, immunocytofluorescence, and scanning electron microscopy. Keywords Blood-brain barrier, Brain endothelial cells, Brain microvessel isolation, Immunocytofluorescence, Pericytes, Western blot

1

Introduction The blood-brain barrier selectively regulates the passage of soluble and potentially toxic molecules from the capillary lumen into the brain parenchyma [1, 2]. Those barrier properties depend on endothelial cells, which interact intimately with pericytes [3– 5]. Brain endothelial cells and pericytes are crucial for the establishment and maintenance of the blood-brain barrier; thus, it is important to study their molecular profile and morphology under physiological and pathological conditions [6–8]. The study of each cellular type and their interactions will improve the understanding of the molecular mechanisms that modulate blood-brain barrier function at the local level. Previous studies show that the blood-brain barrier permeability to circulating molecules is heterogeneous among cerebral regions, according to their metabolic demand, the local release of inflammatory mediators, and the regional differences in astroglial and microglial density [9– 12]. However, the study of the regionalized function of the blood-brain barrier at the level of brain endothelial cells and pericytes is currently infrequent. Here, we describe an easy method to efficiently isolate blood microvessels that contain endothelial cells

37

38

Fernanda Medina-Flores et al.

and pericytes with a high purity, from specific brain regions, such as the cerebral cortex and the hippocampus. This technique is a modification from Tontsch and Bauer [13] in which the cortical and subcortical tissue is used for primary culture of brain microvessels or pericytes. The isolated brain endothelial cells and pericytes obtained using the present method are viable to perform conventional molecular and histological techniques such as Western blot, immunocytofluorescence, and scanning electron microscopy.

2

Materials Prepare fresh solutions with ultrapure water at room temperature. Filter the solutions, and then store them at the corresponding temperature indicated below.

2.1 Solutions for Brain Blood Vessel Isolation

1. 5 PBS solution: Weigh 40.03 g NaCl, 1.01 g KCl, 7.2 g Na2HPO4, and 1.2 g KH2PO4, and transfer them to a 1 L beaker. Add 800 mL ultrapure water to the cylinder, mix with a magnetic stir bar, and adjust pH to 7.4 with a solution of 0.3 M HCl (pH 0.13) or 0.5 M NaOH (pH 13.6). Make up to 1 L with ultrapure water and mix. Store at 4  C. 2. 1 PBS solution: Add 200 mL of 5 PBS solution to the 1 L beaker. Add 600 mL of ultrapure water, mix, and adjust pH to 7.4 with a solution of 0.3 M HCl or 0.5 M NaOH. Make up to 1 L with water; mix with a magnetic stir bar. Filter the solution with sterile syringe filter, pore size 0.2 μm, and store at 4  C. 3. 1% Bovine serum albumin (BSA) solution: Weigh 1 g BSA, add 50 mL filtered 1 PBS solution to the 100 mL graduated beaker, mix with a magnetic stir bar, and adjust pH to 7.4 with a solution of 0.3 M HCl or 0.5 M NaOH. Make up to 100 mL with 1 PBS, and filter it with sterile syringe filter, pore size 0.2 μm. Store at 20  C (see Note 1). 4. Sucrose buffer (SB): 0.32 M sucrose, 3 mM HEPES, and 1% BSA solution. Weigh 10.95 g sucrose and 0.071493 g HEPES, and transfer to 100 mL graduated beaker. Add 50 mL of 1% BSA solution, mix with a magnetic stir bar, and adjust pH to 7.4 with a solution of 0.3 M HCl or 0.5 M NaOH. Make up to 100 mL with 1% BSA solution, mix, and filter it with sterile syringe filter, pore size 0.2 μm. Store at 20  C (see Note 1).

2.2 Solutions for Western Blot

1. Protease inhibitor cocktail: Weigh 2.6298 g NaCl, 2.3634 g Tris–HCl, and 0.5583 g EDTA, and transfer to 15 mL conical tube. Add 3 mL of Triton X-100; mix with a vortex shaker. Add 1% protease inhibitor cocktail, and mix with vortex shaker. Store at 4  C.

Regional Isolation of Brain Microvessels

39

2. 5 Transfer buffer: Glycine 0.9 M, Tris-base 0.15 M, and SDS 0.001 M. Weigh 72 g glycine, 15.15 g tris-base, and 0.5 g SDS, and transfer to 1 L graduated beaker. Add 1 L of deionizer water, and mix with a magnetic stir bar. Store at 4  C. 3. 5 Running buffer: Glycine 0.9 M, Tris-base 0.15 M, and SDS 0.001 M. Weigh 72 g glycine, 15.15 g Tris-base, and 5 g SDS, and transfer to 1 L graduated beaker. Add 1 L of deionizer water, and mix with a magnetic stir bar. Store at 4  C. 4. 5% Blocking solution: Weigh 5 g non-fat dry milk, and transfer to 100 mL graduated beaker. Add 100 mL 1 PBS and mix with a vortex shaker. Use immediately. 5. Washing solution: Weight 1 g non-fat dry milk, and transfer to graduated beaker. Add 100 mL 1 PBS and 100 μL Tween-20 detergent, and mix with a magnetic stir bar. Use immediately or store at 4  C. 6. Avidin-biotin complex solution (Vectastain ABC Elite Standard kit, Vector PK- 6100): In a 15 mL conical tube, add 25 μL reagent A, 25 μL reagent B, and 10 mL 1 PBS; mix with a vortex shaker. Store at 4  C (see Note 2). 2.3 Solutions for Immunocytofluorescence

1. 4% Paraformaldehyde: Weigh 4 g paraformaldehyde, and transfer it to graduated beaker. Add 60 mL warm distilled water, and mix with a magnetic stir bar. Adjust pH to 7.4 with a solution of 0.3 M HCl or 0.5 M NaOH. Make up to 100 mL distilled water, mix, and filter it with sterile syringe filter, pore size 0.2 μm. Store at 4  C. 2. 1 PBS/0.1% Tween-20 detergent: In a 100 mL graduated beaker, add 100 mL 1 PBS and 10 μL Tween-20 detergent, mix with a magnetic stir bar, and use immediately or store at 4  C.

2.4 Solutions for Scanning Electron Microscopy

1. 1% OsO4: Weigh 1 g OsO4 and transfer it to amber glass bottle. Add 100 mL distilled water; mix with a magnetic stir bar until the osmium crystals dissolve. Wrap the cap of the bottle with four or five Parafilm layers. Place it inside a sealed container, and store at 4  C (see Note 3). 2. 4% Paraformaldehyde/2% glutaraldehyde: Weigh 4 g paraformaldehyde, and transfer it to graduated beaker. Add 60 mL warm distilled water, and mix with a magnetic stir bar. Adjust pH to 7.4 with a solution of 0.3 M HCl or 0.5 M NaOH. Later, add 2 mL glutaraldehyde, and make up to 100 mL distilled water, mix, and filter it with sterile syringe filter, pore size 0.2 μm. Store at 4  C. 3. Ethanol solutions: 100% ethanol and distilled water to prepare 30, 40, 50, 70, 80, 90, and 100% ethanol solutions. Mix and store at room temperature.

40

3

Fernanda Medina-Flores et al.

Methods

3.1 Blood Vessel Isolation from Selective Brain Regions

1. Dissection instruments must be cleaned and placed in antiseptic solution diluted in water 1:1000 to sterilize for 1 h at room temperature. 2. Add 1.5 mL 1% BSA solution to each 1.5 mL conical microtube, and store over night at 4  C. This avoids the attachment of microvessels to the walls of the microtubes. 3. Discard the 1% BSA solution from microtubes by decantation. Add 1 mL of SB to the microtube, and maintain it on ice during the experiment. 4. Obtain the head of one euthanized rat, and immediately place it on ice (see Note 4). 5. Use large scissors to remove the skin that is on top of the skull. Next, use dental extracting forceps to remove the skull, begin by carefully introducing the forceps tip on the foramen magnum at the midline level. Carefully lift the skull, and insert again the dental extracting forceps as many times as required performing the lifting movement. You may also use a piglet teeth clipper to cut the skull, beginning at the lateral sides of the foramen magnum. Remove the skull pieces with serrated tweezers, and cut as many times as required. Once fully exposed, carefully remove the brain with medium size serrated tweezers. Cut the residual pia mater with curved iris scissors, and remove it with serrated tweezers. Soak the brain surface with 5 mL ice SB using a plastic Pasteur pipette to eliminate blood residues. 6. Place the brain on filter paper on ice. Next, use iris scissors to cut and obtain the cerebral cortex and hippocampus (see Note 5). 7. Put each brain region on filter paper. Carefully roll the cerebral cortex or hippocampus on the filter paper to eliminate superficial blood vessels, and then transfer the brain tissue to the microtubes with SB solution. 8. Homogenize the tissue in 1 mL ice SB solution using a plastic homogenizer, and pestle for less than 1 min. Make it carefully to avoid the formation of bubbles. 9. Centrifuge the suspension at 1000  g, 4  C for 10 min, and aspirate the supernatant using a 1000 μL pipette. Add 1 mL of ice SB to the pellet, and gently resuspend it with the pipette tip. Centrifuge it again in the same conditions, and discard the supernatant (see Note 6). If there is a layer of myelin that looks like a white layer at the top of the pellet also aspirate it.

Regional Isolation of Brain Microvessels

41

10. Add 1 mL of ice SB, and slowly resuspend the pellet with the pipette tip, and centrifuge at 100  g, 4  C for 45 s. Keep the supernatant, and place it in a new microtube on ice; the supernatant contains the brain blood microvessels. Discard the pellet. 11. Centrifuge the supernatant at 200  g, 4  C for 3 min. Keep only the pellet, as it contains the brain blood vessels, and wash it with 500 μL SB, and then centrifuge it at 200  g, 4  C for 2 min, and discard the supernatant. Rinse the pellet with 500 μL PBS 1, and centrifuge it in the same conditions. Discard the supernatant. The pellet contains the isolated brain blood microvessels. Use the pellet immediately or store at 80  C in 0.6 mL microtubes. 3.2 Protein Expression in Isolated Blood Microvessels from Specific Brain Regions: Western Blot

1. Add to the pellet 250 μL of protease inhibitor cocktail, and gently resuspend it with the pipette tip. Centrifuge the suspension at 19,242  g, 4  C for 10 min. 2. Obtain the supernatant and immediately use it or store it in aliquots at 80  C. 3. Take one aliquot of the supernatant, and perform the protein quantification assay according to the Bradford technique or any other you use in the laboratory. 4. Load 30 μg of total protein in vertical SDS-PAGE 10% polyacrylamide gel. 5. Place the set inside the electrophoresis chamber, and perform the protein migration at 70 V for 30 min, and change to 120 V for 1–2 h, until the bands of the molecular weight marker are set apart from each other. 6. Soak a membrane of polyvinylidene fluoride (PVDF) in 100% methanol for 5 min. Thereafter, place the PVDF membrane in transfer buffer for at least 5 min. 7. Assemble the “sandwich” in the bottom of the cassette with presoaked sponge, filter paper, gel, the PVDF membrane, filter paper, and sponge. Use a blot roller to remove the air bubbles or by gently pressing the “sandwich.” 8. Place the sandwich in a standard transfer wet chamber at 70 V for 60 min in the cold. 9. Carefully remove the PVDF membrane from the cassette, and rinse it with ultrapure water for 5 min, followed by a 100% methanol rinse for 5 min. Use immediately or let it dry and store at 4  C. 10. Block unspecific binding sites by incubating the PVDF membrane with 25 mL blocking solution; place it on a shaker platform for 30 min.

42

Fernanda Medina-Flores et al.

11. Incubate the PVDF membrane overnight with primary antibodies to detect blood-brain barrier proteins (e.g., anti-claudin-5, anti-occludin, anti-ZO-1), pericyte marker (antiPDGFRβ), or other proteins required for your research at 4  C. Before the evaluation of the proteins of interest, it is recommended to incubate the PVDF membrane with antibodies to detect glial and neuronal markers (e.g., GFAP, Iba-1, NeuN) in order to corroborate the purity of the isolated cells. Microvessel samples isolated using this technique have been shown to have a high purity [14]. 12. Discard the primary antibodies, and rinse the PVDF membrane with 5 mL washing solution in constant agitation for 30 min at room temperature. Discard and change the washing solution each for 10 min. 13. Incubate the PVDF membrane with secondary antibodies for 2 h in constant agitation at room temperature. 14. Discard the secondary antibodies, and rinse the PVDF membrane three times with 5 mL ice washing solution for 10 min each at room temperature. 15. Wash the PVDF membrane with 5 mL ice 1 PBS for 10 min at room temperature in agitation. 16. Incubate the PVDF membrane with 5 mL avidin-biotin complex solution for 30 min at room temperature, in constant agitation. 17. Wash the PVDF membrane with 5 mL washing solution for 15 min at room temperature. 18. Reveal the PVDF membrane with chemiluminescence detection system. 19. Acquire the images with a C-DiGit® image generation and analysis system or the equipment of your laboratory. 3.3 Protein Immunoreactivity in Isolated Blood Microvessels from Specific Brain Regions: Immunocytofluorescence and Confocal Microscopy

1. After obtaining the pellet that contains the isolated brain microvessels, add immediately 1 mL of ice 4% paraformaldehyde, and incubate for 1 h at 4  C. 2. Centrifuge at 19,242  g, 4  C for 10 min, and discard the supernatant by decantation. 3. Resuspend and rinse the pellet with 1 mL ice 1 PBS, and then centrifuge at 19,242  g, 4  C for 10 min. Repeat rinses with 1 PBS until the paraformaldehyde odor is eliminated. 4. Add to the pellet of the isolated brain microvessels 1 mL ice 1 PBS. Use a very thin brush to take a small portion of the pellet, and place it gently on a gelatin-coated slide. Slowly distribute the microvessels along the gelatin-coated slide (see Note 7).

Regional Isolation of Brain Microvessels

43

5. Use an optical microscope to check out the distribution of brain microvessels on the gelatin-coated slide (see Note 8). Verify that the microvessels are evenly distributed and separated from each other along the slide. Then, allow the brain microvessels to attach to the slide in a humid chamber for 30 min at room temperature. 6. Slowly add 20 μL 4% paraformaldehyde to the slide, and carefully distribute it throughout the sample to fix the cells, and incubate in humid chamber for 10 min at room temperature. Thereafter, eliminate the paraformaldehyde by slow decantation, and rinse the sample with 20 μL 1 PBS/0.1% Tween-20 (1 PBST). 7. Add on the slide 20 μL blocking serum solution, and distribute it on the sample. Incubate in humid chamber for 15 min at 20  C. Then discard the blocking solution, and rinse the sample once with 20 μL 1 PBST. 8. Incubate the sample with 20 μL primary antibody diluted in 1 PBST for 4 h at 20  C in humid chamber (see Note 9). Thereafter, discard the antibody solution, and carefully wash the sample once with 20 μL 1 PBST. 9. Incubate the isolated brain microvessels with 20 μL fluorescent-labeled secondary antibody for 4 h at 20  C in humid chamber. Discard the antibody solution, and carefully wash the sample once with 20 μL 1 PBST. 10. Eliminate excess solution and add one drop of mounting medium to the slide. Use the forceps to take a coverslip, and slowly place it on the mounting medium, avoiding the formation of bubbles. 11. Apply clear nail polish or glue along the edges to seal the coverslip to the slide. 12. Visualize the isolated brain microvessels using a confocal microscope. 3.4 Morphological Analysis of Pericyte Attachment in Isolated Blood Microvessels from Specific Brain Regions: Scanning Electron Microscopy

1. After obtaining the pellet that contains the isolated brain microvessels, add immediately 1 mL ice 4% paraformaldehyde/2% glutaraldehyde for 24 h at 4  C. 2. Centrifuge at 19,242  g, 4  C for 10 min, and discard the supernatant by decantation. 3. Resuspend and rinse the pellet with 1 mL ice 1 PBS, and then centrifuge at 19,242  g, 4  C for 10 min. Repeat rinses with 1 PBS until the paraformaldehyde-glutaraldehyde odor is eliminated. 4. Add to the pellet of the isolated brain microvessels 1 mL ice 1 PBS. Use a very thin brush to take a small portion of the pellet,

44

Fernanda Medina-Flores et al.

and place it gently on a gelatin-coated coverslip. Slowly distribute the microvessels along the gelatin-coated coverslip. 5. Post-fix with 1% OsO4 for 2 h at room temperature, and then dehydrate in graded ethanol series for 10 min for each one (see Note 10). 6. Use fine tip tweezers to carefully transfer the samples to a microporous capsule with lid. Avoid samples from air drying (see Note 11). 7. Submit samples to the critical point drying procedure of your lab, followed by gold coating. 8. Apply double-sided conductive tape on the surface of the microscope specimen stub, and gently place the dried sample on it (see Note 12). 9. Acquire scanning electron micrographs at 13 kV with a scanning electron microscope.

4

Notes 1. The volume of the solution depends on the number of samples you will work with; 100 mL of 1% BSA solution and sucrose buffer are sufficient to process two brain regions from six rat brains. 2. The avidin-biotin complex solution must be prepared and stored at 4  C 30 min before using it. 3. Osmium tetroxide is extremely toxic; use coat, gloves, and goggles to prepare it in a lab extraction hood. 4. The euthanasia method does not modify the viability of brain microvessels. Animals may be euthanized by anesthesia overdose (e.g., with sodium pentobarbital) or by any other accepted method for rats. Consider obtaining the tissue quickly as possible, and process it immediately. 5. To dissect the cerebral cortex, use iris scissors, and carefully cut horizontally to the brain surface a piece of 1.5 cm long and 3 mm depth for each hemisphere. To dissect the hippocampus, remove the debris from the cerebral cortex with medium serrated tweezers using a gentle movement from midline to the lateral side of each hemisphere. The debris are easily cleared due to the presence of the lateral ventricles underneath the cerebral cortex. Once exposed, cut a piece of 2–3 mm thick and 1 cm long of the hippocampus with iris scissors using a lateral toward a midline movement. Repeat for the other brain hemisphere. 6. Carefully aspire the supernatant because some microvessels can be attached to the wall of the microtube.

Regional Isolation of Brain Microvessels

45

7. Distribute and perfectly separate the small portion of the pellet on the gelatin-coated slide. Avoid leaving large fragments of the pellet on the slide because the microvessels will be seen as tangles under the microscope ensuing poor detection of tight junction protein expression. 8. Observing the isolated brain microvessels with the optical microscope will let you know if the microvessels are attached to the gelatin-coated slide. If the brain microvessels are floating and constantly moving on the slide, they will be lost during sample processing. In this case, verify the viability of the gelatin-coated slides, and repeat step 4, and then observe the sample again under the microscope. 9. The antibody dilution will be prepared according to the one normally used in your laboratory. 10. Ethanol must be slowly added down the wall of the multiple well plate to avoid direct pressure on the sample. Use a different plastic Pasteur pipette to slowly add and remove the ethanol because small portions of the pellet could remain suspended. Avoid the Pasteur pipette tip to touch the sample surface. 11. After the sample is placed inside the microporous specimen capsule and perfectly closed, transfer it to a polyethylene specimen cup with 100 mL 100% ethanol. If the microporous capsule floats on the surface, it means there is air inside it, and the sample must be removed and fixed again. 12. Use thin-tip scissors to distribute the sample over the stub, and ensure that it is completely adhered. References 1. Chow BW, Gu C (2015) The molecular constituents of the blood-brain barrier. Trends Neurosci 38:598–608. https://doi.org/10. 1016/j.tins.2015.08.003 2. Keaney J, Campbell M (2015) The dynamic blood-brain barrier. FEBS J 282 (21):4067–4079. https://doi.org/10.1111/ febs.13412 3. Armulik A, Abramsson A, Betsholtz C (2005) Endothelial/pericyte interactions. Circ Res 97 (6):512–523. https://doi.org/10.1161/01. RES.0000182903.16652.d7 4. Bell RD, Winkler EA, Sagare AP et al (2010) Pericytes control key neurovascular functions and neuronal phenotype in the adult brain and during brain aging. Neuron 68(3):409–427. https://doi.org/10.1016/j.neuron.2010.09. 043

5. Rustenhoven J, Jansson D, Smyth LC et al (2017) Brain Pericytes as mediators of Neuroinflammation. Trends Pharmacol Sci 38 (3):291–304. https://doi.org/10.1016/j. tips.2016.12.001 6. Hamilton NB (2010) Pericyte-mediated regulation of capillary diameter: a component of neurovascular coupling in health and disease. Front Neuroenerg 2:2. https://doi.org/10. 3389/fnene.2010.00005 7. Sweeney MD, Sagare AP, Zlokovic BV (2018) Blood–brain barrier breakdown in Alzheimer disease and other neurodegenerative disorders. Nat Rev Neurosci 14(3):133–150. https:// doi.org/10.1038/nrneurol.2017.188 8. Winkler EA, Bell RD, Zlokovic BV (2011) Central nervous system pericytes in health and disease. Nat Neurosci 14(11):1398–1405. https://doi.org/10.1038/nn.2946

46

Fernanda Medina-Flores et al.

9. Banks WA, Kastin AJ (1998) Differential permeability of the blood-brain barrier to two pancreatic peptides: insulin and amylin. Peptides 19(8):883–889. https://doi.org/10. 1016/S0196-9781(98)00018-7 10. Hurtado-Alvarado G, Vela´zquezMoctezuma J, Go´mez-Gonza´lez B (2017) Chronic sleep restriction disrupts interendothelial junctions in the hippocampus and increases blood-brain barrier permeability: chronic sleep restriction disrupts interendothelial junctions. J Microsc 268(1):28–38. https://doi.org/10.1111/jmi.12583 11. Moinuddin A, Morley JE, Banks WA (2000) Regional variations in the transport of interleukin-1α across the blood-brain barrier in ICR and aging SAMP8 mice. Neuroimmunomodulation 8:165–170. https://doi.org/10.1159/ 000054814

˜ or R, Kuennecke B, Ozmen L et al 12. Villasen (2017) Region-specific permeability of the blood–brain barrier upon pericyte loss. J Cerebr Blood F Met 37(12):3683–3694. https:// doi.org/10.1177/0271678X17697340 13. Tontsch U, Bauer HC (1989) Isolation, characterization, and long-term cultivation of porcine and murine cerebral capillary endothelial cells. Microvasc Res 37(2):148–161. https:// doi.org/10.1016/0026-2862(89)90034-4 14. Medina-Flores F, Hurtado-Alvarado G, Contis-Montes de Oca A, Lo´pez-Cervantes S, Ko¨nigsberg M, Deli MA, Go´mez-Gonza´lez B (2020) Sleep loss disrupts pericyte-endothelial cell interactions impairing blood-brain barrier function. Brain Behav Immun. https://doi. org/10.1016/j.bbi.2020.05.077

Methods in Molecular Biology (2021) 2367: 47–72 DOI 10.1007/7651_2020_311 © Springer Science+Business Media New York 2020 Published online: 14 August 2020

Estimating Brain Permeability Using In Vitro Blood-Brain Barrier Models Saeideh Nozohouri, Behnam Noorani, Abraham Al-Ahmad, and Thomas J. Abbruscato Abstract The blood-brain barrier (BBB) is a vital biological interface that regulates transfer of different molecules between blood and brain and, therefore, maintains the homeostatic environment of the CNS. In order to perform high-throughput screening of therapeutics in drug discovery, specific properties of the BBB are investigated within in vitro BBB platforms. In this chapter, we detail the process and steps for the iPSC to BMEC and astrocyte differentiation as well as TEER and permeability measurement in Transwell platform of in vitro BBB model. Also, advanced microfluidic iPSCs-derived BMECs on chip and permeability measurement within this model have been elucidated. Keywords BBB, iPSC, Endothelial cells, Astrocytes, Co-culture model, Transwell insert, In vitro permeability, Microfluidic chips

1

Introduction Although CNS disorders are the major causes of mortality and morbidity worldwide, therapeutic options for these diseases, compared to any other organ system, are currently limited [1]. One of the main obstacles in drug development of neurotherapeutics is the presence of the blood-brain barrier (BBB) that restricts access to the brain for more than 98% of small molecules and virtually all macromolecules [2]. The BBB maintains the homeostatic environment of the CNS and plays an essential role in the mass transfer between the circulatory system and brain tissue [3–5]. It consists of specialized endothelial cells with a basal lamina that supports the abluminal surface of the endothelium along with other supporting cells, such as pericytes, astrocytes, and neurons [6]. The inner space of cerebral capillaries is covered by brain microvascular endothelial cells (BMECs), which are different from endothelial cells in other organs in their protein expression and function. Particularly, they are connected by complex tight-junction proteins that effectively

Saeideh Nozohouri and Behnam Noorani contributed equally to this work.

47

48

Saeideh Nozohouri et al.

seal the paracellular pathway and limit the passage of molecules [6]. These cells also contain specific transporters on the luminal and abluminal membranes of the capillaries, which allow for exchange of essential nutrients between the brain and the blood transcellularly, along with eliminating waste and harmful substances [6]. Therefore, BMECs are the primary gatekeepers for the transport and efflux of nutrients, metabolites, and neurotoxins. To evaluate the role of BBB in regulating the permeability and transport of different molecules in healthy and disease conditions, numerous in vivo and in vitro models have been utilized [4, 7, 8]. Considering the complexity of the BBB function and structure, in vivo models provide the most reliable measurements of drug permeability and remain the gold standard for this purpose [9, 10]. However, animal-based in vivo techniques are high cost, low throughput, and labor-intensive [10]. Given the demand for high-throughput methods in order to screen numerous compounds, there has been a pronounced interest in in vitro models. In vitro BBB models have been used widely in modern times because of their improved reproducibility and low cost; therefore, they can be used in performing high-throughput screening studies as well as their ability to complete mechanistic studies that examine regulatory mechanisms at the molecular level. Different cerebrovascular ECs (primary, immortalized, and PSC-) derived from a variety of sources can be used in a multitude of experimental setups [11–18]. However, primary cells are not easy to purify, and they lose necessary BBB phenotypes quickly. Furthermore, immortalized BMECs do not always provide good in vitro barrier properties because of their low transendothelial electrical resistance (TEER) values, which makes them unsuitable for permeability studies [19, 20]. Current concerns and disadvantages related to the primary or immortalized cells derived from animals and humans (Table 1) have led to examination of human pluripotent stem cells in order to obtain BMECs [17, 25]. These endothelial cells contain attractive characteristics such as self-renewal capability, which creates an unlimited source of cells and improved barrier and efflux transporters properties [17]. These cells also provide physiological TEER values that result in low paracellular permeability. Moreover, other cells of NVU, including astrocytes, pericytes, and neurons, can be differentiated from human induced pluripotent stem cells (iPSCs), which enables creating a fully human BBB model [28– 31]. Therefore, this alternative cell line generates in vitro data that correlates well with in vivo studies. An efficient in vitro BBB model is crucial for facilitating the rapid screening of neurotherapeutics’ permeability. The selected in vitro model should mimic functionalities of the BBB by expressing important BBB efflux proteins and having lower permeability in comparison with other in vitro models. The chosen model should also demonstrate the most significant and related features

Estimating Brain Permeability Using In Vitro Blood-Brain Barrier Models

49

Table 1 Sources of cells used to obtain BMEC for in vitro studies Cell type

Cell origin

Advantages

Primary

Mouse, rat, human, bovine, porcine

l

l

Closest similarity to the BBB phenotype in vivo Demonstrate excellent characteristics of the BBB at early passages

Disadvantages l

l l

l

l

l

l

Immortalized

Pluripotent stem cellderived

Human colon Adenocarcinoma Epithelial (Caco-2) l Mouse BMEC (BEnd.3/ BEnd.5) l Rat BMEC (RBE4) l Human BMEC (hCMEC/D3 and hBMEC)

l

Human

l

l

l

l

l l

l l

Commercially available Possibility of modification to express human proteins Stable over higher passage numbers Low cost Easy growth

l

Very high TEER Self-renewal capacity Low passive permeability

l

l l

l

References

Difficult to obtain pure [19–21] cells Low yield of isolation Require the sacrifice of animals Senesce of cells after few passages Human origin is not suitable for highthroughput studies Time-consuming and costly to isolate Losing BBB characteristics over passages Barrier function is negatively affected by the immortalization process Low TEER values Leaky

[22–24]

Needs differentiation and characterization High cost

[17, 25–27]

of the BBB based on the particular goal of the specific investigation and the availability of resources [32]. The main challenge with the development of effective in vitro models is to recognize and utilize pure cell types in order to achieve high TEER values with low permeability. Up to the early 1980s, the different non-endothelial components of the neurovascular unit (NVU) have been mostly documented as non-essential to the barrier function, providing only a scaffold function to the blood-brain barrier. However, the pioneer in vivo work of Janzer and Raff in 1987 [33] on chick embryos, in addition to the in vitro study of Arthur [34], highlighted the

50

Saeideh Nozohouri et al.

importance of astrocytes in the induction of the barrier phenotype in both CNS and non-CNS endothelial cells. In both studies, the authors documented the ability of astrocytes to induce barrier tightness when co-cultured (or implanted at close proximity of a vascular tree) with endothelial cells. As of today, the nature of such factors still remains mostly unidentified, although certain studies identified angiopoietin-1 (Ang-1) [35] or retinoic acid (RA) [36]. Interestingly, astrocytes are the last cell type of the NVU to arise during development [37, 38]. During development, brain endothelial cells invade a primitive neural tissue formed by neuronal stem cell (NSC)/progenitor cells (NPCs) and differentiating neurons [39], followed by the recruitment of pericytes [11] and ultimately astrocytes through the formation of end-feet processes [37, 38]. Astrocytes and neurons share a common cell lineage origin, coming from a neural crest origin [40]. Such differentiating neuroepithelium will give rise to a neuronal stem cell (NSC) population lining the subventricular zone (SVZ), the inner part of the neural tube in the primitive encephalon. Such NSC present in the SVZ differentiates into two cell types: neuroblasts (which will mature and form neurons) and the radial glia (RG). Astrocytes originate from the RG and chronologically engulf and populate the BBB as the last cell type. Differentiation of human pluripotent stem cells (hPSCs) into neurons and glial cells (including astrocytes) follows a temporal differentiation pattern similar to in vivo. Stem-cell derived astrocytes are obtained from neuronal progenitor cells (NPC), by maintenance and expansion of NPCs grown as neurospheres for over 60 days. Such NPCs are generated by differentiation of hPSCs into neuroepithelium which are visible in tissue culture plates as formation of “rosettes.” Such rosettes can be enzymatically dissociated and resuspended into neurospheres which further differentiate into NSC and NPCs using defined differentiation medium. As of today, there is no unified differentiation protocol for astrocytes used as co-cultures, as different protocols have been described in the literature. Such protocols display different iteration whether neurospheres are obtained from NPCs isolated from primary cultures [41, 42] or from hPSC-derived neurospheres [43]. The most common in vitro platforms for creating BBB models are generated using Transwells. These models can be developed by using human or murine brain-derived endothelial cells, which are seeded on a permeable membrane that is coated with an extracellular matrix [18]. The cells are grown in the form of a single monolayer or in co-cultivation with astrocytes to create a co-culture model [18]. The endothelial cells are grown in the luminal compartment of the Transwell, and other cells are cultured in the abluminal side of the Transwell insert (Fig. 1). The endothelial cells can either be human stem cell-derived brain endothelial cells and immortalized

Estimating Brain Permeability Using In Vitro Blood-Brain Barrier Models

51

Fig. 1 Schematic representative of Transwell apparatus and BBB co-culture model (Created by Biorender.com)

human cell lines such as hCMEC/D3 cells [44] or animal-derived immortalized or primary cell lines [18, 45], which are most commonly used cells. Some advantages of the Transwell® system include scalability, ease of use, and the possibility of quantifying barrier integrity by measuring TEER. Drawbacks of this platform are the lack of flow and shear stress, three-dimensional structure, and the presence of membrane between cells, which limits the contact of different cell types within the model. The performance of this model should be evaluated by quantitative assessment of barrier function, which can be performed through measuring TEER and permeability of different markers. If this permeability is in acceptable physiological levels, then the BBB in vitro model can be used for testing drug candidates. The permeability measurement of the in vitro models requires the use of suitable markers. Various BBB permeability markers are currently being used for in vitro studies, including sodium fluorescein, sucrose, dextran, etc. (Table 2). Among the drug-like low molecular weight markers, sodium fluorescein and radiolabeled sucrose (e.g., [14C] sucrose) are used widely in in vitro experiments with high TEER values [17]. Sucrose is considered the most accurate marker because it is metabolically stable, is uncharged, and does not bind to proteins; however, the radioactive isotope of

52

Saeideh Nozohouri et al.

Table 2 Different paracellular markers for in vitro permeability studies Name of marker

Molecular weight Advantage(s)

Sodium fluorescein

376 Da

– Easy detection method – Only weakly binds to proteins – Freely diffusible – Detectable in very low concentrations – Inexpensive – Non-radioactive – Nontoxic

– Slight interaction with BBB transporters

[14C] sucrose

342 Da

– No interaction with proteins – Metabolically stable – Uncharged

– – – – –

[13C12] sucrose

354 Da

– Requires LC-MS/MS device for – Non-radioactive detection – Sensitive and specific method of detection – Stable isotope – Metabolically inert – No interaction with proteins – Uncharged

[47]

Dextran

4–70 kDa

– Not suitable for small molecules – Can be used for a permeability prediction broad range of molecular weights – Stability issue (FITC can detach from dextran and misleads for – Easy detection higher permeability) method – Can be conjugated – May be toxic at high concentrations with FITC or biotin

[46]

Disadvantage(s)

Reference (s) [46]

[47] Radioactive Contains lipophilic impurities Over-time degradation High cost Nonspecific measurement of total radioactivity with a scintillation counter – Requires radioactive license

sucrose has shown to contain some lipophilic impurities, and performing experiments with this marker needs radioactive license and permission [47]. Recently, [13C12] sucrose as a superior marker has been introduced, which is non-radioactive and can be quantified by a sensitive and highly specific LC-MS/MS technique [48, 49]. This isotope may be selected as the most accurate marker for in vivo experiments, but there is not any report about the in vitro application of this tracer yet. Sodium fluorescein, which was introduced as a small molecular weight tracer (MW 376), has

Estimating Brain Permeability Using In Vitro Blood-Brain Barrier Models

53

Fig. 2 General steps of iPSC differentiation to BMECs and astrocytes (Created by Biorender.com)

been widely used in a variety of models for the investigation of BBB permeability [50]. Fluorescein, available as its salt form, is a very small molecule, freely diffusible, nontoxic, and detectable at very low concentrations [50]. Owing to its small size, sodium fluorescein may cross the BBB much more readily than larger molecular weight tracers such as dextran and is more sensitive to changes in BBB integrity, while using large molecular weight markers (>3 kDa) may not accurately show the integrity of the BBB model. Therefore, sodium fluorescein can be one of the most useful tracers due to being inexpensive, non-radioactive, and easy to detect for in vitro experiments. In the following protocol, we have described a permeability experiment by using sodium fluorescein; however, other permeability markers (mentioned in Table 2) can also be utilized based on the availability of the materials and the aims of the study. Different sources of cells are currently being used for the in vitro permeability study of BBB. Based on the availability of the materials and the purpose of the study, proper cell lines should be selected. Due to the aforementioned properties of iPSC-derived endothelial cells, they can be an appropriate cell source for BBB drug screening investigations. Therefore, we often utilize the iPSC line as an in vitro model for clarifying permeability measurements of a drug candidate. Also, in this chapter, we will describe the differentiation protocol implemented by Patel and colleagues in their isogeneic model of the human BBB (Fig. 2) [51]. This protocol allows for the differentiation of astrocytes and neurons using adherent cells rather than floating neurospheres, which eases the differentiation protocol. iPSC-derived astrocytes showed expression of several cell markers considered typical for human astrocytes (GFAP+, S100β+, Nestin-), were able to induce barrier properties in iPSC-derived BMECs monolayers, and responded to environmental injury (e.g., hypoxia/ischemia) [52] similar to primary astrocytes monocultures

54

Saeideh Nozohouri et al.

or co-cultured with BMECs [53–56]. We will also describe one of the most common platforms of in vitro models, “Transwell” filters, in this protocol. Furthermore, recent advanced in vitro models of BBB and their consecutive results are briefly discussed.

2

Materials This protocol was adapted from Al Ahmad and Lippmann publications [17, 26].

2.1 Materials for Differentiation of iPSC to hBMECs

2.2 Preparation of Aliquots and Mediums

l

iPSC line that exhibits minimal spontaneous differentiation. We recommend using the IMR90-4 hiPSC line since this cell line has extensively been validated.

l

Essential 8 Medium (500 mL, Fisher Scientific).

l

Matrigel (Corning hESC-Qualified matrix, 5 mL, Fisher Scientific).

l

Gibco™ Knockout™ Serum Replacement (500 mL, Fisher Scientific).

l

DMEM/F12, 15 mM HEPES (500 mL, Fisher Scientific).

l

ReLSR (Enzyme-free human ES and iPS cell selection and passaging reagent, Fisher Scientific).

l

ROCK inhibitor Y27632 (R&D Systems).

l

Gibco™ MEM Non-Essential Amino Acids Solution (100 mL, Fisher Scientific).

l

Gibco™ GlutaMAX™ Supplement (100 mL, Fisher Scientific).

l

Mercaptoethanol cell culture grade (Sigma-Aldrich).

l

Human endothelial serum-free medium (hESFM, 500 mL, Fisher Scientific).

l

Platelet-poor plasma-derived serum (PDS) (Fisher Scientific).

l

Basic human fibroblast growth factor recombinant (bFGF, Peprotech).

l

Retinoic acid (RA) (10 mM Sigma-Aldrich).

l

Extracellular matrix (ECM) coating solution including collagen IV (Sigma-Aldrich) and fibronectin (Sigma-Aldrich).

l

Accutase (Fisher Scientific).

l

DMSO (Sigma-Aldrich).

l

500 mL and 100 mL Fisherbrand™ Disposable PES Filter Units

l

Tissue culture 6-well plates (Corning).

l

Matrigel aliquots: Corning Matrigel hESC-qualified matrix should be reconstituted, aliquoted, and stored as recommended in the product guidelines for use.

Estimating Brain Permeability Using In Vitro Blood-Brain Barrier Models

55

1. Cool microcentrifuge tubes and pipet tips in the
20  C freezer the day before. 2. The aliquot (dilution factor) volume is typically between 270 and 350 μL and is calculated for each lot based on the protein concentration. 3. Prepare around 16 aliquots at the time, quickly label, and put them immediately in the
80  C (please note that the Matrigel can transform to gel quickly at 22–35  C). 4. Prepare a Working Solution of Corning Matrigel hESCqualified matrix; add one aliquot in 25 mL of DMEM/ F12 and plate 6-well plates with 1 mL/well. 5. Allow the Matrigel to coat for at least 1 h. Plates can be kept in the incubator for up to 5 days (see Note 1). l

Rock inhibitor Y-27632 aliquots 1. Stock solutions of 10 mM are prepared (50 μL aliquots) in sterile ddH2O and stored at
20  C.

l

KnockOut serum aliquots 1. Allow the KnockOut serum to thaw at 4  C overnight. (please note that the knockout serum should not thaw on warm water). 2. Aliquot the knockout serum into 50 mL conical tubes and freeze at
20  C.

l

PDS aliquots 1. Thaw overnight at 4  C and centrifuge the plasma in 50 mL conical tubes at 18,800  g for 10 min at 4  C. 2. Recover the supernatant and avoid the precipitate. 3. Aliquot in 1 mL tubes and store at
20  C.

l

BFGF aliquots 1. Dissolve 100 μg of powder in sterile ddH2O and aliquot 20 μg/mL (1000). 2. Final concentration of BFGF in medium should be 20 ng/ mL, so 100 μL aliquots (20 μg/mL) should be prepared for 100 mL medium.

l

RA aliquots 1. Reconstitute retinoic acid in DMSO at a concentration of 10 mmol/mL. 2. Aliquot in 100 μL tubes and store at
20  C. 3. Final concentration should be 10 μmol/mL.

56

Saeideh Nozohouri et al. l

Accutase aliquots 1. Thaw the Accutase solution at 4  C overnight, aliquot into 6 mL tubes, and store at
20  C. 2. Accutase is stable at 4  C for 72 h.

l

Collagen solution (5 mL) 1. Prepare 15 mM acetic solution in sterile water. 2. Add 5 mL acetic acid solution in collagen powder (final concentration 1 mg/mL). 3. Allow collagen to dissolve on the bench for 30 min and store at 4  C.

l

Preparation of mediums 1. Unconditioned medium (UM) (500 mL). 2. 100 mL knockout (KO) serum aliquots (20%). KO serum aliquots should be allowed to thaw overnight at 4  C. (The serum should not thaw using warm tap water or water bath). 3. 5 mL Non-Essential Amino Acid from the stock solution (1%). 4. 2.5 mL GlutaMAX from stock solution (0.5%). 5. 7.8 μL of b-mercaptoethanol from stock solution (0.1 mM). 6. Fill the filter unit to 500 mL, filter the medium, and keep at 4  C for up to 4 weeks.

l

EC++ (100 mL) 1. Fill the filter unit to 100 mL with hESFM. 2. 100 μL bFGF aliquot (20 ng/mL). 3. 100 μL RA aliquot (10 μmol/mL). 4. 1 mL PDS (1%). 5. Filter the medium and keep at 4  C for up to 3 weeks.

l

EC

(100 mL) 1. Fill the filter unit to 100 mL with hESFM. 2. 1 mL PDS (1%). 3. Filter the medium and keep at 4  C for up to 3 weeks.

l

1 Collagen/fibronectin mixture for insert coating (1 mL) 1. 400 μL collagen. 2. 100 μL fibronectin. 3. 500 μL sterile ddH2O. 4. For coating plates and other surfaces, you can dilute five times with ddH2O (5 mL total volume) (see Note 2).

Estimating Brain Permeability Using In Vitro Blood-Brain Barrier Models

57

2.3 Materials for Differentiation of iPSC to Astrocytes

l

PSC Neural Induction Medium Kit (NIM, 500 mL, Life Tech).

l

StemPro™ hESC serum-free medium (hESC SFM, 500 mL, Life Tech).

2.3.1 Reagents

l

GlutaMAX I Supplement (100 mL, Life Tech).

l

Human recombinant brain-derived neurotrophic factor (BDNF, Peprotech).

l

Human recombinant glia-derived neurotrophic factor (GDNF, Peprotech).

l

Laminin from EHS murine sarcoma basement membrane (Sigma-Aldrich).

l

Poly-D-lysine hydrobromide (PDL, Sigma-Aldrich).

l

B-27™ Plus Neuronal Culture System (500 mL, Life Tech).

l

CultureOne™ Supplement 100 (100 mL, Life Tech).

l

hESC-qualified Matrigel (Corning).

l

Astrocytes Medium Kit (500 mL, Life Tech).

l

Neural Induction Medium (NIM)—50 mL solution

2.3.2 Mediums

1. Mix 1 mL of Neural Induction Supplement (50) with 50 mL of neurobasal medium. (Both media are from the Neural Induction Medium Kit.) l

Neural Differentiation Medium (NDM)—100 mL solution 1. 7.2 mL of BSA solution at 25% (from hESC SFM Kit). 2. 2 mL of hESC supplement (from hESC SFM Kit). 3. 1 μg BDNF (40 μL from 25 μg/mL stock solution). 4. 1 μg GDNF (10 μL from 100 μg/mL stock solution). 5. 1 mL GlutaMAX I (Life Technologies). 6. Add DMEM/F12 from the hESC SFM kit to 100 mL.

l

Astrocytes Maturation Medium (AMM)—100 mL solution 1. 1 mL OneShot FBS provided with the Astrocyte Medium Kit. 2. 1 mL N2 supplement provided with the Astrocyte Medium Kit. 3. Complete to 100 mL with DMEM from the Astrocyte Medium Kit (4.5 g/L glucose).

2.4 Materials for Permeability and TEER Measurement

l

Sodium fluorescein (Sigma-Aldrich).

l

Electrode probes (World Precision Instruments).

l

Millicell® ERS Voltohmmeter or EVOM Voltohmmeter.

l

96-well clear-bottom black microplate (Fisher Scientific).

l

Synergy™ Mx Microplate reader (Bio-Tek).

58

3

Saeideh Nozohouri et al.

Methods

3.1 Differentiation of iPSCs to hBMECs

l

Seeding iPSC line in 6-well plates 1. Thaw the iPSC line vial. 2. Quickly resuspend the cell pellet in 5 mL of E8 medium in a 15 mL centrifuge tube. 3. Spin down cells at 170  g for 5 min. 4. Prepare E8 containing 10 μM Y27632 and resuspend the cell pellets with it. 5. Seed the iPSC cells with E8 medium containing 10 μM Y27632 on Matrigel-coated well (6-well plate). 6. Change the medium 24 h after seeding, switch to a regular E8 medium (2 mL each well), and feed the wells daily until days 3–4. 7. Passage the iPSCs at a 1:6 ratio into freshly coated plates when 60–80% confluence. – Passage ratios are empirically determined based on the growth rate of the iPSC line.

l

Passaging iPSC lines 1. When the cells are 60–80% confluent (days 3–4), remove the E8 medium and add 0.5 mL of ReLeSR per well and keep it at 37  C for 2 min. 2. Aspirate the ReLeSR from your passaging well and add 6 mL of E8 in the well, and wash out the well three times by aspiration/reflux; clumps should be visible with naked eyes. 3. Aspirate the Matrigel from the coated wells, and add 1 m of suspended cells to each well. 4. Add 1 mL of E8 medium to each well to have a total volume of 2 mL/well.

l

Differentiation of iPSCs to BMEC 1. iPSC lines are ready to differentiate when the confluency reaches to 60–80%. 2. Add 0.5 mL Accutase into each well. Incubate at 37  C for 10 min. 3. Neutralize Accutase by adding 2 mL of E8 medium. Collect the iPSCs from each well, and resuspend cells in a conical tube. 4. Spin down the cells via centrifugation at 170  g for 5 min.

Estimating Brain Permeability Using In Vitro Blood-Brain Barrier Models

59

5. Aspirate supernatant and resuspend cells in 1 mL of E8 medium. 6. Count the cell density with an automated cell counter or hemocytometer. 7. Plate cells at a density of 10,000 cells/cm2 (100,000 cells per well of 6-well plate) in E8 medium containing 10 μM Y27632 in Matrigel-coated plate (2 mL E8 medium each well). 8. 24 h after seeding, change the medium with 2 mL of regular E8 for each well daily until 3 days. 9. Initiate the differentiation by replacing the iPSC maintenance medium with the UM medium. 10. Replace medium with fresh UM every day (e.g., 2 mL per well, 6-well plate). 11. After 6 days in UM, change medium to EC++. Do not change the medium for 48 h. 12. 48 h after switching medium to EC++, which would be day 8 of differentiation, cells are ready for further experiments and seeding on Transwell filters. 13. At day 8 of differentiation, dissociate cells with Accutase. Add 0.5 mL Accutase into each well. Incubate at 37  C for 30 min. 14. Neutralize the Accutase treatment with 2 mL UM, gently collect cells by spraying across each well two to three times using a P1000 pipet, and transfer the cell suspension to conical tubes. 15. Centrifuge the cell suspension at 170  g for 5 min. 16. Aspirate supernatant and resuspend the cell pellet in a proper volume of EC++ (0.5 mL/Transwell filter). – Coat 12-well Transwell filters 4–24 h before the cells are ready for seeding with 200 μL of collagen/fibronectin solution (ECM solution), and keep them in the incubator at 37  C. 17. Remove the ECM solution from the coated Transwell filters by aspiration. 18. Seed the cells at a density of 106 cells/cm2 in 0.5 mL EC++ per well of Transwell filters. Add 1.5 mL of EC++ to each basolateral chamber. 19. 24 h after subculture, change medium from EC++ to EC

to induce barrier formation (see Note 3).

60

Saeideh Nozohouri et al.

3.2 Differentiation of iPSC into Astrocytes

1. Initiate the differentiation of undifferentiated iPSCs by >80% confluency stage by replacing the iPSC maintenance medium with 1 NIM. 2. Replace differentiating iPSCs with fresh NIM medium daily for 10 days. Differentiation efficacy can be checked by performing immunostaining against PAX6 (neuroepithelium cell marker). 3. At day 11 of differentiation, dissociate cells into single cells using Accutase (R) for 10 min. Centrifuge cells and resuspend cells in NIM. Count cell density in resuspension. 4. Seed Matrigel-coated plates with cells at a density of 105 cells/ cm2, in presence of NIM supplemented with 10 μM Y-27632 for 24 h. 5. 24 h after reseeding cells, replace NIM with NDM. 6. Feed cells daily with NDM for 5 days. 7. Dissociate cells with Accutase for 10 min and centrifuge cells. Cells can be frozen at this stage as NPCs and resumed for differentiation as needed. 8. Seed cells on freshly Matrigel-coated plates (ideally 12-well or 24-well plates) at a cell density of 2.5  104 cells/cm2 in presence of NDM for 24 h. 9. 24 h after seeding, replace the NDM medium with AMM. 10. Replace AMM every 2 days for 2–3 weeks, to synchronize astrocytes to BMECs differentiation. 11. At day 8 of BMECs differentiation, add the Transwell insert into the astrocyte well immediately before BMECs seeding. 12. Seed BMECs as previously described and maintain these cells in EC+/+. 13. 24 h after seeding, replace cell medium in both compartments. Add EC
/
in the apical (top) chamber, and add AMM in the basolateral (bottom) chamber. 14. 48 h after seeding, cell co-cultures are ready for assessing the barrier function or for performing experiments.

3.3 Permeability and TEER Measurement

The TEER should be measured 24 h after EC

or 48 h after subculture of cells on the Transwell filter.

3.3.1 For Measuring the TEER

1. Sterilize chopsticks by using 70% ethanol, and let the ethanol evaporate from the surface of the electrodes. 2. Measure the TEER by placing the electrodes carefully on each Transwell without damaging the membrane. Please note that the shorter electrode should be in the upper channel and the longer electrode should be placed at the bottom of the filter. Repeat the measurement for three times on each filter.

Estimating Brain Permeability Using In Vitro Blood-Brain Barrier Models

61

3. Subtract each measured resistance from the resistance of an empty filter in order to obtain the TEER value associated with the endothelial cells’ monolayer. Then, multiply each value by the surface area of the used filter to calculate the final value in Ω  cm2. 4. First, measure the empty coated filter and then filters containing cells. 5. We recommend choosing samples with TEER values higher than 1,000 Ω  cm2 for any permeability study (see Notes 4 and 5). 3.3.2 Permeability Measurement

1. Prepare transport buffer in advance with distilled water and 0.12 M NaCl, 25 mM NaHCO3, 3 mM KCl, 2 mM MgSO4, 2 mM CaCl2, 0.4 mM K2HPO4, 1 mM HEPES, and 0.1% human platelet-poor derived serum. 2. Dilute sodium fluorescein to create a stock solution of 10 mM in distilled water. The stock solution can be stored in an opaque box at 4  C. Prepare a working solution of 10 μM sodium fluorescein by diluting the stock solution in the transport buffer. 3. Using the transport buffer, prepare standard curve concentrations of sodium fluorescein from 10 μM working solution. First dilute the 10 μM solution 12.5 times to get a concentration of 800 nM, and then prepare 8 dilution series of 800 nM, 400 nM, 200 nM, 100 nM, 50 nM, 25 nM, 12.5 nM, and 0 nm in 200 μL. We recommend taking the same volume (200 μL) from Transwells while sampling at each time point. Place each dilution and sample into a well of 96-well plate. 2. Remove the medium from both chambers of each filter, and add 1.5 mL of medium to the basolateral chamber and then add 0.5 mL of sodium fluorescein solution to the donor chamber. 3. Take 50 μL from the 10 μM stock solution, and dilute it 20 times to be in the range of standard curve (200 μL final volume) and transfer it to the 96-well plate. We will use this value as the donor chamber concentration in the calculations. 4. Collect 200 μL at 15, 30, 60, 90, and 120 min from the basolateral chamber, and transfer the collected samples to the 96-well plate and then add 200 μL fresh media to each basolateral compartment to replace the removed volume return the plate to the incubator. 5. Make sure to cover the 96-well plate with aluminum foil after each collection. This is because sodium fluorescein is light sensitive.

62

Saeideh Nozohouri et al.

6. After 2 h, measure the fluorescence of each well using a fluorescent microplate reader with excitation and emission wavelengths of 485 and 520 nm. 7. Create a standard curve by plotting fluorescence versus concentration. 8. Calculate the concentration of each time point by using the calibration curve (see Notes 6 and 7). 9. Calculate the cleared volume of each time point from Eq. 1: Cleared Volume ¼ ðC ðabluminalÞ  V ðabluminalÞ=C ðluminalÞ ð1Þ l

C abluminal refers to measured concentration in abluminal compartment at a given sampling time point.

l

V abluminal refers to the volume of abluminal compartment.

l

C luminal refers to concentration in luminal compartment.

10. Plot the calculated cleared volume versus time for each sample as well as the empty filters. 11. Apply linear regression and obtain the slope of each filter. 12. The slope of the regression line corresponds to the “clearance” or permeability-surface area product (“PS” or “PA”) of the Transwell system. 13. To obtain the permeability coefficient, P (Eq. 2), the clearance value has to be divided by the filter area (e.g., 1.12 cm2 per well for a 12-well Transwell filter): P ¼ PS=S

ð2Þ

Here, P is permeability coefficient, PS is permeability-surface area product or clearance, and S is the surface of the insert in cm2. 14. Check the units when calculating P. The unit of P is cm/s; therefore, convert the units of min to second and μL to mm3 and then to cm3. Please note that the most common cause of misinterpretation of the results in this assay is inconsistent units. 15. The permeability of the cell monolayer alone can be extracted from the Transwell permeability (total permeability) by correcting for the permeability of an empty filter based on Eq. 3: 1=Pcells ¼ 1=Ptotal
1=Pfilter ðsee Notes 8
11Þ

ð3Þ

Estimating Brain Permeability Using In Vitro Blood-Brain Barrier Models

3.4 Advanced Microfluidic iPSCsDerived hBMECs on Chips

63

The main goal of in vitro models of BBB is to acquire an environment outside a living organism to evaluate the physiological and pathological responses to any stimuli and provide a model for drug development and discovery. Currently, there are no viable alternatives for in vivo experiments in the preclinical phase of drug development for obtaining good estimates of permeability characteristics and distribution of small and large molecular weight agents to the brain tissue. Therefore, progress in understanding the physiology, in particular the transport mechanisms of the BBB by advanced in vitro models, is highly desirable. Various in vitro BBB models have been reported and characterized in terms of barrier properties, expression profile of BBB-related proteins, and applicability for pharmacological studies [10, 57, 58]. The organ-on-a-chip as a new class of in vitro models has recently been introduced by recent advanced technologies such as 3D printing and soft lithographic techniques [59]. In this model, human cells can be used in a microenvironment that reproduces a tissue similar to in vivo conditions. This more physiologically related microenvironment can reduce the gap between traditional in vitro models and in vivo models with human physiology by providing 3D structures and dynamic conditions. Moreover, these platforms can be suitable for the modeling of biological barriers, especially the BBB, because the flow in the microchannel mimics the blood flow, and the hydrogel can serve as the extracellular matrix for different cell types. These new advanced models can compensate the lack of physiologically related conditions of the Transwell system, such as exposure to shear stress (mechanical stimuli) and the complex 3D environment of the BBB [10]. Since the application of iPSC-derived brain endothelial cells in permeability studies showed promising results due to high TEER values, expression of tight-junction proteins, and low permeability, these ECs have recently been applied in microfluidic devices and have shown the permeability results close to in vivo studies [31, 60–63]. The permeability outcomes and the key features of previous BBB-on-chip models in which iPSC-derived brain endothelial cells were used have been summarized in Table 3. The permeability coefficients of a compound (cm/s) obtained for passive transport can be compared with other permeability coefficient values of the same compound in vivo or in other platforms, including Transwell system. For instance, Svendsen’s group showed a correlation (R2 ¼ 0.96) between permeability values of different molecular weight dextran (3, 20, and 70 kDa) across BBB-chips and brain uptake of these markers in rodents [64]. The apparent permeability of 4 kDa FITC-Dextran was approximately 8  10
7 cm/s, while the apparent permeability of 4 kDa FITC-Dextran on post-capillary venules of rats was 0.92  10
6 cm/s [64]. Moreover, Searson’s group measured the permeability of Lucifer yellow on iPSCderived human BMECs tissue-engineered microvessels, and the

BMECs derived from human iPSCs BMECs derived from human iPSCs

Gelatin hydrogel with Human collagen 800 μm diameter type IV and channel (fabricated by fibronectin cylindrical template rod)

Human collagen type IV and fibronectin

Human collagen type IV and fibronectin

Human collagen type IV and fibronectin

Collagen matrix with 150 μm cylindrical diameter channel

PDMS with two closely opposed, parallel microchannels separated by a porous membrane (1  1 mm and 1  0.2 mm)

Collagen matrix with 150 μm cylindrical diameter channel

BMECs derived from human iPSCs

BMECs derived from human iPSCs

BMECs derived from human iPSCs

Biological coating

Endothelial cell type

Human collagen type IV and fibronectin

PDMS with two closely opposed, parallel microchannels separated by a porous membrane (1  1 mm and 1  0.2 mm)

Material and structure of microvessels

NA

Primary human pericytes and astrocytes

NA

NA

Primary human pericytes and astrocytes

Additional cells in co-culture

Lucifer yellow: 3–4  10
7 Dextran: below detection limit

Sodium fluorescein: 1–3  10
6 Dextran: 4.3  10
8 Albumin 1  10
10

0.8–1  10
7

Lucifer yellow and 10 kDa dextran

24,000 not normalized

NA

NA

1,500

TEER (Ω  cm2)

Lucifer yellow ¼2–3  10
7 NA The permeability of 10 kDa dextran in microvessels was below the detection limit

Fluorescent8.9, 1.1, and 0.24  10
8 labeled for 3, 10, and 70 kDa, dextran tracers respectively (3, 10, or 70 kDa)

Lucifer yellow, Alexa Flour10 kDa dextran

Sodium fluorescein, 3 kDa FITCdextran, albumin

Fluorescentlabeled dextran tracer (3 kDa)

Barrier Permeability measurement permeability tracer (cm/s)

Table 3 Summary of current iPSC-derived BBB organ-on-chips and their key features

4

6

1

0.3

2.4

[62]

[63]

[65]

[61]

[64]

Shear stress (dyn/cm2) References

64 Saeideh Nozohouri et al.

Fibronectin

PDMS containing fibrin gel with self-organized channels with less than 100 μm diameter

PDMS with parallel Human collagen microchannels with type IV and 300 μm  160 μm with fibronectin polycarbonate insert with 04 μm pore size

Human collagen type IV and fibronectin

Collagen matrix with 150 μm cylindrical diameter channel

BMECs derived from human iPSCs

BMECs derived from human iPSCs

BMECs derived from human iPSCs

Rat primary astrocytes

Primary human pericytes and astrocytes

Human iPSCpericyte

NA

2000–4,400

10 kDa dextran: 2.2 10
7 NA 40 kDa dextran: 8.9  10
8

Lucifer yellow ¼ 4  10
7

4, 20 and 70 KDa 4 kDa dextran ¼10
7 20 and 70 kDa ¼ 10
8 FITC-dextran

40 and 10 kDa FTIC-dextran

Lucifer yellow

0.25

NA

4

[60]

[66]

[31]

Estimating Brain Permeability Using In Vitro Blood-Brain Barrier Models 65

66

Saeideh Nozohouri et al.

reported permeability was low (2–3  10
7 cm/s) which was close to reported value of post-capillary venules in rats (1–2  10
7 cm/s) [62]. Based on the results from Table 3, these models form much tighter and more durable barriers that can mimic the in vivo integrity due to the presence of shear stress, microenvironment, and cellcell interaction; moreover, the in vitro and in vivo correlation can be obtained with these advanced models. In spite of recapitulating the neurovascular unit, providing a single protocol for permeability measurement in these models is difficult due to the possibility of various design and structural inconsistencies that alter quantification. In general, the transport of small and large molecular weight agents can be evaluated and quantified in real time across microfluidic chips via fluorescent microscope techniques. In this technique, the molecule of interest is labeled with a fluorescent tag and perfused through the constructed vascular channel with a particular flow rate. Real-time images are obtained and analyzed to investigate the rate of molecule permeability into the acceptor tissue chamber from the vascular channel. The permeability is calculated based on the rate of change in fluorescent intensity in the tissue compartments by Eq. 4 [62, 67]: p¼

1 V dI I 0 S dt

ð4Þ

where I0 is the maximum fluorescence intensity of the vascular channel containing endothelial cells, (dI/dt) is the rate of increase in fluorescent intensity as solute enters into the tissue compartment, and V/S is the ratio of apical volume to the surface area. In detail, the desired concentration of fluorescent-tagged molecule is prepared in transport media and loaded into the reservoir (it can be syringe or reservoir tubes based on the pump). Then, the perfused tubing containing the fluorescent molecule is connected to the chip. Please note that the cell-loaded microfluidic chip should be placed onto the stage of a fluorescence microscope (inverted microscope recommended). Then, the specified flow rate begins, and immediately the process of capturing images starts. Each microscope software has the ability to acquire images automatically, or it can be done manually at the defined time point. Based on the size of the marker and flow rate, images are captured every minute for at least 1 h (total frames and time of the process of imaging depend on how fast the fluorescent molecule can enter the tissue compartment). The apical and basolateral channels of the chip should be taken in one image so the objective and exposure view should be adjusted accordingly. To prevent photo bleaching, the fluorescence shutter should be closed between image acquisitions.

Estimating Brain Permeability Using In Vitro Blood-Brain Barrier Models

67

Afterwards, the intensity profile is obtained by using Image J (NIH). The image sequences are converted to AVI format and then imported to Image J. Also, vascular channel geometry, based on the region of interest, is obtained by using rectangle section tool. Then, the mean gray value of the stack is analyzed using plugins and stacks section of the software and then the stack is measured by image J and the data is exported from the results window to Excel for further studies. Please note that the mean gray value can be obtained by selecting mean gray value of the particular file in Set measurement in the Analyze section of software. Intensity of vascular channel achieves from this data (I0). The same processes are performed for tissue compartment to obtain the intensity of that compartment. Then, time is calculated by multiplying each frame number by total second of the experiment (e.g., 1 h ¼ 3,600 s) and dividing by the number of the frames (total frames ¼ 60). Plot I microvessel channel vs time in an Excel scatter plot. The average gray value intensity of the plateau area will be I0 in Eq. (4). Next, find the slope of Itissue vs. time. Only use data in the time range specified by the I0 average above. Plot Itissue data in the y-axis and time in the x-axis. Display the trend line with its equation on the graph to determine the slope. The slope is equal to dI/dt in Eq. (4) [67]. Therefore, the obtained endothelial permeability coefficient can be compared with other permeability coefficients of the same analyte in vivo or other platforms including Transwell system. A potential drawback is the requirement of using a fluorescent tag which can alter the chemistry, size, and biological activity of the test molecule.

4

Notes 1. If Matrigel hESC-qualified matrix is not thawed properly, clumps might be created in the solution. Keep Matrigel vial on ice all the time while handling. 2. Please note that in the recent publication of Lippmann’s group, B27 (200 dilution, Thermo Fisher Scientific) has been used instead of PDS in endothelial cell medium preparation and resulted in hBMECs of equal fidelity compared to previously reported method with higher TEER value. They showed that replacement of serum with fully defined components (B27) yields BBB endothelium with TEER in the range of 2,000–8,000 cm2 across multiple iPSC lines, with appropriate marker expression and active transporters [68]. The use of a fully defined medium vastly improves the consistency of differentiation, and co-culture of BBB endothelium with iPSCderived astrocytes produces a robust in vitro neurovascular

68

Saeideh Nozohouri et al.

model [68]. Therefore, another option in preparing EC++ and EC

is substituting PDS with B27 (200 dilution). 3. Changing medium from EC++ to EC

(EC++ depleted of bFGF and RA), TEER reaches the maximum after 24 h and often lowers drastically over 2–3 days. Therefore, it is recommended to measure permeability and other experiments between 24 and 48 h after switching EC++. 4. A minimum of three filters seeded with hBMECs are required to perform the permeability study of any marker or drug. In addition, to obtain mass transfer resistance, two coated Transwell filters without the cells should be used per experiment. 5. Start the process of permeability measurement after measuring the TEER and ensuring the monolayers are still intact. 6. Please note that the fluorescence of each sample should be calculated based on the volumetric difference of each sample to obtain the correct concentration. This is because the fluorescence from each sample is for 1.5 mL volume in the basolateral chamber while the standard curve was created with 200 μL volumes. 7. Keep in mind to consider the fact that removing fluorescein from the bottom chamber after each time point results in a decrease in fluorescence. To correct each time point and to account for the removed fraction, you should add the concentration from the previous time point multiplied by 0.2/1.5. 8. The time points of this assay can be determined based on the size of the paracellular marker. For large molecular weight markers, the experiment duration can be more than 2 h, and the sampling time points may start after 60 min. 9. Eliminate bubbles before reading fluorescence using a needle. 10. Make sure the reading value of TEER machine is stable before recording since sometimes it takes time for the electrode to show the correct number. 11. We recommend using iPSC cell line with passage numbers less than 50 for any permeability study due to the decrease in TEER values. References 1. Lozano R, Naghavi M, Foreman K et al (2012) Global and regional mortality from 235 causes of death for 20 age groups in 1990 and 2010: a systematic analysis for the Global Burden of Disease Study 2010. Lancet 380 (9859):2095–2128. https://doi.org/10. 1016/S0140-6736(12)61728-0

2. Pardridge WM (2007) Blood-brain barrier delivery. Drug Discov Today 12(1–2):54–61. https://doi.org/10.1016/j.drudis.2006.10. 013 3. Paulson OB (2002) Blood-brain barrier, brain metabolism and cerebral blood flow. Eur Neuropsychopharmacol 12(6):495–501

Estimating Brain Permeability Using In Vitro Blood-Brain Barrier Models 4. Cecchelli R, Berezowski V, Lundquist S et al (2007) Modelling of the blood-brain barrier in drug discovery and development. Nat Rev Drug Discov 6(8):650–661. https://doi.org/ 10.1038/nrd2368 5. Nozohouri S, Sifat AE, Vaidya B, Abbruscato TJ (2020) Novel approaches for the delivery of therapeutics in ischemic stroke. Drug Discov Today 25(3):535–551. https://doi.org/10. 1016/j.drudis.2020.01.007 6. Abbott NJ, Patabendige AA, Dolman DE et al (2010) Structure and function of the blood-brain barrier. Neurobiol Dis 37 (1):13–25. https://doi.org/10.1016/j.nbd. 2009.07.030 7. Hawkins BT, Davis TP (2005) The bloodbrain barrier/neurovascular unit in health and disease. Pharmacol Rev 57(2):173–185. https://doi.org/10.1124/pr.57.2.4 8. Wilhelm I, Krizbai IA (2014) In vitro models of the blood-brain barrier for the study of drug delivery to the brain. Mol Pharm 11 (7):1949–1963. https://doi.org/10.1021/ mp500046f 9. Wolff A, Antfolk M, Brodin B, Tenje M (2015) In vitro blood-brain barrier models-an overview of established models and new microfluidic approaches. J Pharm Sci 104 (9):2727–2746. https://doi.org/10.1002/ jps.24329 10. Saunders NR, Dziegielewska KM, Mollgard K, Habgood MD (2015) Markers for blood-brain barrier integrity: how appropriate is Evans blue in the twenty-first century and what are the alternatives? Front Neurosci 9:385. https:// doi.org/10.3389/fnins.2015.00385 11. Daneman R, Zhou L, Kebede AA, Barres BA (2010) Pericytes are required for blood-brain barrier integrity during embryogenesis. Nature 468(7323):562–566. https://doi.org/10. 1038/nature09513 12. Faria A, Pestana D, Teixeira D et al (2010) Flavonoid transport across RBE4 cells: a blood-brain barrier model. Cell Mol Biol Lett 15(2):234–241. https://doi.org/10.2478/ s11658-010-0006-4 13. Hatherell K, Couraud PO, Romero IA et al (2011) Development of a three-dimensional, all-human in vitro model of the blood-brain barrier using mono-, co-, and tri-cultivation Transwell models. J Neurosci Methods 199 (2):223–229. https://doi.org/10.1016/j. jneumeth.2011.05.012 14. Abbott NJ, Dolman DE, Drndarski S, Fredriksson SM (2012) An improved in vitro blood-brain barrier model: rat brain endothelial cells co-cultured with astrocytes. Methods

69

Mol Biol 814:415–430. https://doi.org/10. 1007/978-1-61779-452-0_28 15. Paolinelli R, Corada M, Ferrarini L et al (2013) Wnt activation of immortalized brain endothelial cells as a tool for generating a standardized model of the blood brain barrier in vitro. PLoS One 8(8):e70233. https://doi.org/10.1371/ journal.pone.0070233 16. Helms HC, Brodin B (2014) Generation of primary cultures of bovine brain endothelial cells and setup of cocultures with rat astrocytes. Methods Mol Biol 1135:365–382. https:// doi.org/10.1007/978-1-4939-0320-7_30 17. Lippmann ES, Al-Ahmad A, Azarin SM et al (2014) A retinoic acid-enhanced, multicellular human blood-brain barrier model derived from stem cell sources. Sci Rep 4:4160. https://doi. org/10.1038/srep04160 18. Helms HC, Abbott NJ, Burek M et al (2016) In vitro models of the blood-brain barrier: an overview of commonly used brain endothelial cell culture models and guidelines for their use. J Cereb Blood Flow Metab 36(5):862–890. https://doi.org/10.1177/ 0271678X16630991 19. Bernas MJ, Cardoso FL, Daley SK et al (2010) Establishment of primary cultures of human brain microvascular endothelial cells to provide an in vitro cellular model of the blood-brain barrier. Nat Protoc 5(7):1265–1272. https:// doi.org/10.1038/nprot.2010.76 20. Kaisar MA, Sajja RK, Prasad S et al (2017) New experimental models of the blood-brain barrier for CNS drug discovery. Expert Opin Drug Discovery 12(1):89–103. https://doi.org/10. 1080/17460441.2017.1253676 21. Thomsen LB, Burkhart A, Moos T (2015) A triple culture model of the blood-brain barrier using porcine brain endothelial cells, astrocytes and Pericytes. PLoS One 10(8):e0134765. https://doi.org/10.1371/journal.pone. 0134765 22. Weksler BB, Subileau EA, Perriere N et al (2005) Blood-brain barrier-specific properties of a human adult brain endothelial cell line. FASEB J 19(13):1872–1874. https://doi. org/10.1096/fj.04-3458fje 23. Vu K, Weksler B, Romero I et al (2009) Immortalized human brain endothelial cell line HCMEC/D3 as a model of the bloodbrain barrier facilitates in vitro studies of central nervous system infection by Cryptococcus neoformans. Eukaryot Cell 8(11):1803–1807. https://doi.org/10.1128/EC.00240-09 24. Eigenmann DE, Xue G, Kim KS et al (2013) Comparative study of four immortalized human brain capillary endothelial cell lines,

70

Saeideh Nozohouri et al.

hCMEC/D3, hBMEC, TY10, and BB19, and optimization of culture conditions, for an in vitro blood-brain barrier model for drug permeability studies. Fluids Barriers CNS 10 (1):33. https://doi.org/10.1186/20458118-10-33 25. Lippmann ES, Azarin SM, Kay JE et al (2012) Derivation of blood-brain barrier endothelial cells from human pluripotent stem cells. Nat Biotechnol 30(8):783–791. https://doi.org/ 10.1038/nbt.2247 26. Patel R, Alahmad AJ (2016) Growth-factor reduced Matrigel source influences stem cell derived brain microvascular endothelial cell barrier properties. Fluids Barriers CNS 13:6. https://doi.org/10.1186/s12987-016-0030-5 27. Katt ME, Xu ZS, Gerecht S, Searson PC (2016) Human brain microvascular endothelial cells derived from the BC1 iPS cell line exhibit a blood-brain barrier phenotype. Plos One 11 (4):e0152105. https://doi.org/10.1371/jour nal.pone.0152105 28. Shaltouki A, Peng J, Liu Q et al (2013) Efficient generation of astrocytes from human pluripotent stem cells in defined conditions. Stem Cells 31(5):941–952. https://doi.org/10. 1002/stem.1334 29. Pei Y, Peng J, Behl M et al (2016) Comparative neurotoxicity screening in human iPSCderived neural stem cells, neurons and astrocytes. Brain Res 1638(Pt A):57–73. https:// doi.org/10.1016/j.brainres.2015.07.048 30. Albekairi TH, Vaidya B, Patel R et al (2019) Brain delivery of a potent opioid receptor agonist, Biphalin during ischemic stroke: role of organic anion transporting polypeptide (OATP). Pharmaceutics 11(9). https://doi. org/10.3390/pharmaceutics11090467 31. Jamieson JJ, Linville RM, Ding YY et al (2019) Role of iPSC-derived pericytes on barrier function of iPSC-derived brain microvascular endothelial cells in 2D and 3D. Fluids Barriers CNS 16(1):15. https://doi.org/10.1186/s12987019-0136-7 32. Jamieson JJ, Searson PC, Gerecht S (2017) Engineering the human blood-brain barrier in vitro. J Biol Eng 11:37. https://doi.org/ 10.1186/s13036-017-0076-1 33. Janzer RC, Raff MC (1987) Astrocytes induce blood-brain barrier properties in endothelial cells. Nature 325(6101):253–257. https:// doi.org/10.1038/325253a0 34. Arthur FE, Shivers RR, Bowman PD (1987) Astrocyte-mediated induction of tight junctions in brain capillary endothelium: an efficient in vitro model. Brain Res 433 (1):155–159

35. Lee SW, Kim WJ, Choi YK et al (2003) SSeCKS regulates angiogenesis and tight junction formation in blood-brain barrier. Nat Med 9 (7):900–906. https://doi.org/10.1038/ nm889 36. Mizee MR, Wooldrik D, Lakeman KA et al (2013) Retinoic acid induces blood-brain barrier development. J Neurosci 33 (4):1660–1671. https://doi.org/10.1523/J NEUROSCI.1338-12.2013 37. Zerlin M, Goldman JE (1997) Interactions between glial progenitors and blood vessels during early postnatal corticogenesis: blood vessel contact represents an early stage of astrocyte differentiation. J Comp Neurol 387 (4):537–546. https://doi.org/10.1002/( SICI)1096-9861(19971103)387:43.0.CO;2-3. [pii] 38. Zerlin M, Levison SW, Goldman JE (1995) Early patterns of migration, morphogenesis, and intermediate filament expression of subventricular zone cells in the postnatal rat forebrain. J Neurosci 15(11):7238–7249 39. Lippmann ES, Al-Ahmad A, Palecek SP, Shusta EV (2013) Modeling the blood-brain barrier using stem cell sources. Fluids Barriers CNS 10 (1):2. https://doi.org/10.1186/2045-811810-2 40. Akdemir ES, Huang AY, Deneen B (2020) Astrocytogenesis: where, when, and how. F1000Res 9. https://doi.org/10.12688/ f1000research.22405.1 41. Lippmann ES, Weidenfeller C, Svendsen CN, Shusta EV (2011) Blood-brain barrier modeling with co-cultured neural progenitor cellderived astrocytes and neurons. J Neurochem 119(3):507–520. https://doi.org/10.1111/j. 1471-4159.2011.07434.x 42. Weidenfeller C, Svendsen CN, Shusta EV (2007) Differentiating embryonic neural progenitor cells induce blood-brain barrier properties. J Neurochem 101(2):555–565. https:// doi.org/10.1111/j.1471-4159.2006. 04394.x 43. Canfield SG, Stebbins MJ, Morales BS et al (2017) An isogenic blood-brain barrier model comprising brain endothelial cells, astrocytes, and neurons derived from human induced pluripotent stem cells. J Neurochem 140 (6):874–888. https://doi.org/10.1111/jnc. 13923 44. Weksler B, Subileau E, Perriere N et al (2005) Blood-brain barrier-specific properties of a human adult brain endothelial cell line. FASEB J 19(13):1872–1874 45. Nakagawa S, Deli MA, Kawaguchi H et al (2009) A new blood–brain barrier model

Estimating Brain Permeability Using In Vitro Blood-Brain Barrier Models using primary rat brain endothelial cells, pericytes and astrocytes. Neurochem Int 54 (3–4):253–263 46. Grifno GN, Farrell AM, Linville RM et al (2019) Tissue-engineered blood-brain barrier models via directed differentiation of human induced pluripotent stem cells. Sci Rep 9 (1):13957. https://doi.org/10.1038/ s41598-019-50193-1 47. Miah MK, Chowdhury EA, Bickel U, Mehvar R (2017) Evaluation of [(14)C] and [(13)C] sucrose as blood-brain barrier permeability markers. J Pharm Sci 106(6):1659–1669. https://doi.org/10.1016/j.xphs.2017.02. 011 48. Alqahtani F, Chowdhury EA, Bhattacharya R et al (2018) Brain uptake of [13C] and [14C] sucrose quantified by microdialysis and whole tissue analysis in mice. Drug Metab Dispos 46 (11):1514–1518. https://doi.org/10.1124/ dmd.118.082909 49. Chowdhury EA, Alqahtani F, Bhattacharya R et al (2018) Simultaneous UPLC-MS/MS analysis of two stable isotope labeled versions of sucrose in mouse plasma and brain samples as markers of blood-brain barrier permeability and brain vascular space. J Chromatogr B Analyt Technol Biomed Life Sci 1073:19–26. https://doi.org/10.1016/j.jchromb.2017. 12.007 50. Kaya M, Ahishali B (2011) Assessment of permeability in barrier type of endothelium in brain using tracers: Evans blue, sodium fluorescein, and horseradish peroxidase. Methods Mol Biol 763:369–382. https://doi.org/10.1007/ 978-1-61779-191-8_25 51. Patel R, Page S, Al-Ahmad AJ (2017) Isogenic blood-brain barrier models based on patientderived stem cells display inter-individual differences in cell maturation and functionality. J Neurochem 142(1):74–88. https://doi.org/ 10.1111/jnc.14040 52. Page S, Raut S, Al-Ahmad A (2019) Oxygenglucose deprivation/reoxygenation-induced barrier disruption at the human blood-brain barrier is partially mediated through the HIF-1 pathway. Neuromolecular Med. https://doi.org/10.1007/s12017-01908531-z 53. Al Ahmad A, Gassmann M, Ogunshola OO (2012) Involvement of oxidative stress in hypoxia-induced blood-brain barrier breakdown. Microvasc Res 84(2):222–225. https://doi.org/10.1016/j.mvr.2012.05.008 54. Al Ahmad A, Taboada CB, Gassmann M, Ogunshola OO (2011) Astrocytes and pericytes differentially modulate blood-brain barrier characteristics during development and

71

hypoxic insult. J Cereb Blood Flow Metab 31 (2):693–705. https://doi.org/10.1038/ jcbfm.2010.148 55. Al Ahmad A, Gassmann M, Ogunshola OO (2009) Maintaining blood-brain barrier integrity: pericytes perform better than astrocytes during prolonged oxygen deprivation. J Cell Physiol 218(3):612–622. https://doi.org/10. 1002/jcp.21638 56. Schmid-Brunclik N, Burgi-Taboada C, Antoniou X et al (2008) Astrocyte responses to injury: VEGF simultaneously modulates cell death and proliferation. Am J Physiol Regul Integr Comp Physiol 295(3):R864–R873. https://doi.org/10.1152/ajpregu.00536. 2007 57. Phan DT, Bender RHF, Andrejecsk JW et al (2017) Blood-brain barrier-on-a-chip: microphysiological systems that capture the complexity of the blood-central nervous system interface. Exp Biol Med (Maywood) 242 (17):1669–1678. https://doi.org/10.1177/ 1535370217694100 58. Bhalerao A, Sivandzade F, Archie SR et al (2020) In vitro modeling of the neurovascular unit: advances in the field. Fluids Barriers CNS 17(1):22. https://doi.org/10.1186/s12987020-00183-7 59. Zheng F, Fu F, Cheng Y et al (2016) Organon-a-Chip Systems: microengineering to biomimic living systems. Small 12 (17):2253–2282. https://doi.org/10.1002/ smll.201503208 60. Wang YI, Abaci HE, Shuler ML (2017) Microfluidic blood-brain barrier model provides in vivo-like barrier properties for drug permeability screening. Biotechnol Bioeng 114 (1):184–194. https://doi.org/10.1002/bit. 26045 61. Faley SL, Neal EH, Wang JX et al (2019) iPSCderived brain endothelium exhibits stable, long-term barrier function in perfused hydrogel scaffolds. Stem Cell Reports 12 (3):474–487. https://doi.org/10.1016/j. stemcr.2019.01.009 62. Linville RM, DeStefano JG, Sklar MB et al (2019) Human iPSC-derived blood-brain barrier microvessels: validation of barrier function and endothelial cell behavior. Biomaterials 190-191:24–37. https://doi.org/10.1016/j. biomaterials.2018.10.023 63. Park TE, Mustafaoglu N, Herland A et al (2019) Hypoxia-enhanced blood-brain barrier Chip recapitulates human barrier function and shuttling of drugs and antibodies. Nat Commun 10(1):2621. https://doi.org/10.1038/ s41467-019-10588-0

72

Saeideh Nozohouri et al.

64. Vatine GD, Barrile R, Workman MJ et al (2019) Human iPSC-derived blood-brain barrier chips enable disease modeling and personalized medicine applications. Cell Stem Cell 24(6):995–1005.e6. https://doi.org/10. 1016/j.stem.2019.05.011 65. Campisi M, Shin Y, Osaki T et al (2018) 3D self-organized microvascular model of the human blood-brain barrier with endothelial cells, pericytes and astrocytes. Biomaterials 180:117–129. https://doi.org/10.1016/j. biomaterials.2018.07.014 66. van der Helm MW, van der Meer AD, Eijkel JC et al (2016) Microfluidic organ-on-chip

technology for blood-brain barrier research. Tissue Barriers 4(1):e1142493. https://doi. org/10.1080/21688370.2016.1142493 67. Prabhakarpandian B, Shen MC, Nichols JB et al (2013) SyM-BBB: a microfluidic blood brain barrier model. Lab Chip 13 (6):1093–1101. https://doi.org/10.1039/ c2lc41208j 68. Neal EH, Marinelli NA, Shi Y et al (2019) A simplified, fully defined differentiation scheme for producing blood-brain barrier endothelial cells from human iPSCs. Stem Cell Reports 12 (6):1380–1388. https://doi.org/10.1016/j. stemcr.2019.05.008

Methods in Molecular Biology (2021) 2367: 73–85 DOI 10.1007/7651_2020_344 © Springer Science+Business Media, LLC 2021 Published online: 09 March 2021

In Vitro Human Blood-Brain Barrier Model for Drug Permeability Testing Ece Bayir and Aylin Sendemir Abstract Blood-brain barrier (BBB), although very important for protection of brain from major neurotoxins, negatively affects the treatment of central nervous system diseases by limiting the passage of neuropharmaceuticals from blood to the brain. Thus, researchers have to investigate the passage of the produced drug molecules through the BBB before they are introduced to the market. Although these experiments have been traditionally performed on experimental animals, drug permeability tests are now carried out mostly by in vitro BBB models due to ethical problems, differences between species, and expensive and troublesome in vivo test procedures. In this method, we explain how to model and characterize a realistic in vitro BBB model using human derived cells and perform a drug permeability test using this model. Key words Astrocytes, Blood-brain barrier, Brain microvascular endothelial cells, Brain microvascular pericytes, Modeling barrier tissues, Permeability, Tight junctions

1

Introduction Vascular endothelial cells, which cover the interior surface of the blood vessels, are located in all types of blood vessels, from large arteries and veins to the smallest capillaries. The main task of these cells is to regulate the transmission of macromolecules in the bloodstream to the surrounding tissue [1]. Vascular endothelial cells regulate substance permeability through tight junctions (TJs) and adherens junctions (AJs). TJs, located in the apical side of the cells, are multiprotein complexes that act as a fence, which prevents mixing of membrane lipids between the apical and basolateral membranes, and limits the passage of molecules and ions between cells. Although AJs, which are not as tight as TJs, are located in the basolateral side of the cells, they support TJs in limiting the substance permeability by joining the neighboring cells together. The blood-brain barrier (BBB), one of the systems where TJs and AJs are seen most frequently, acts as a dynamic barrier and protects the brain from toxic substances in the bloodstream [2]. BBB mainly consists of brain microvascular endothelial cells (BMECs), a continuous basal lamina, astrocytes, pericytes, and neighboring neurons. BMECs form the vascular structure in the brain and play the

73

74

Ece Bayir and Aylin Sendemir

most important role in BBB. BMECs have very different characteristics than peripheral endothelial cells, including continuous basement membrane, lack of fenestrae structure, low number of pinocytic vesicles, high number of mitochondria, and high transendothelial electrical resistance (TEER) due to their high number of TJs [3, 4]. Astrocytes located between neurons and BMECs play a critical role in BBB by covering a large part of the vascular surface with their cytoplasmic extensions called end-feet. Many astrocytederived factors, such as glial-derived neurotrophic factor (GDNF), basic fibroblast growth factor (bFGF), angiopoietin-1 (ang-1), and transforming growth factor-β (TGF-β), have been reported to induce BBB characteristics of BMECs [5]. Pericytes are mesenchymal cells that cover approximately 30% of the capillary structure formed by BMECs in the abluminal part of the BBB and surrounded by a basal lamina [6]. They play a crucial role in the regulation of angiogenesis and the maintenance of BMECs by growth factors that they secrete, such as TGF-β, platelet-derived growth factor (PDGF), ang-1 and ang-2, and synaptophysin-1 phosphate [7]. BBB and blood-spinal fluid barrier appear as the biggest obstacles for the therapeutics that can be used in the treatment of central nervous system (CNS) diseases to reach damaged neurons [8]. Low molecular weight molecules, such as nicotine, CO2, and O2, with high lipid solubility can cross the BBB through passive diffusion. Substance transport through BMECs depends on molecular structure, size, solubility, affinity, blood flow rate, and expression of carrier proteins. Drugs that are not lipophilic have a molecular weight of more than 400 Da and that form more than eight hydrogen bonds cannot pass BBB through passive diffusion [9]. The development of drugs that can cross the BBB in the treatment of diseases affecting the CNS is challenging due to the highly selective permeability and protected nature of BBB, and only 10% of the developed drugs can enter the clinical trial process [10]. Since most of the large molecules, such as peptides, recombinant proteins, monoclonal antibodies, and ribonucleic acid interference (RNAi) based drugs, developed for use in the treatment of CNS diseases in recent years cannot pass BBB, there has been a considerable increase in the number of studies on regulating drug formulations and overcoming BBB problem through endogenous transport systems [11]. BBB is in the interest of many different disciplines, such as neurology, physiology, pharmacology, and medicine. BBB models are of great importance for studying the behavior of BBB in pathological conditions, such as neurodegenerative diseases, brain tumors, and infection, as well as in the investigation of the mechanisms followed during the transition of neuropharmaceuticals from blood to the brain. CNS-targeted drug development studies begin with library scans of specific molecules against the target and

In Vitro Human Blood-Brain Barrier Model for Drug Permeability Testing

75

continue with the analysis of the molecule’s absorption, distribution, metabolism, and excretion (ADME) properties [12]. ADME studies are usually performed in rodent laboratory animals and later extrapolated to humans; however, during the extrapolation, misleading results can be obtained due to the differences between species [13]. Differences have been shown between rodents and humans in the expression of carrier proteins, receptors, and tightbinding proteins, and therefore BBB permeability [14–16]. Due to the differences in laboratory animals’ and human BBB systems for efficient pharmaceutical investigations, as well as the high cost and long experimental periods needed for animal studies, realistic in vitro models must be constructed using human derived CNS cells. This chapter describes an experimental in vitro BBB model we have developed [17] using a triple co-culture system on standard inserts employing primary human cells (brain microvascular endothelial cells, brain microvascular pericytes and astrocytes) that can be used for testing drug permeability.

2

Materials All chemicals must be suitable for mammalian cell culture. The storage conditions of the solutions should be considered.

2.1 Triple Co-culture Growth Medium Preparation

1. Class II Biosafety Cabinet. 2. Pipetting aid. 3. Micropipettes, 1–10 μL, 10–100 μL, and 100–1000 μL. 4. Dulbecco’s Modified Eagle’s Medium (DMEM), High Glucose (see Note 1). 5. Fetal bovine serum (FBS), sterile, heat inactivated. 6. L-Glutamine, sterile, 200 mM. 7. Sodium Pyruvate, sterile, 100 mM. 8. Gentamycin solution, 50 mg/mL in sterile type 1 ultrapure water (UPW). 9. Putrescine dihydrochloride. 10. Insulin solution, sterile, 1.7 mM. 11. Heparin. 12. Bovine serum albumin (BSA). 13. Hydrocortisone. 14. Epidermal growth factor (EGF). 15. Progesterone. 16. Sodium selenite. 17. Basic fibroblast growth factor (bFGF).

76

Ece Bayir and Aylin Sendemir

18. Sterile bottle for medium preparation. 19. Individually wrapped sterile disposable pipettes. 20. Ethanol, 70%, in spray bottle. 21. Absorbent paper tissues. 2.2

Cell Culture

1. Class II Biosafety Cabinet. 2. Incubator, 37
C, 5% CO2, 95% humidified. 3. Inverted microscope. 4. Refrigerated centrifuge. 5. Pipetting aid. 6. Hemocytometer or electronic cell counter. 7. Human primary brain microvascular endothelial cells (BMECs). 8. Human primary brain microvascular pericytes. 9. Human primary brain astrocytes. 10. Triple co-culture growth medium (see Subheading 2.1). 11. Ca2+ and Mg2+ free phosphate-buffered saline (PBS, 1): 0.2 g/L KCl, 0.2 g/L KH2PO4, 8 g/L NaCl, 2.2 g/L Na2HPO4.7H2O in type 1 UPW, pH 7.4. (Sterilized by filtering through a syringe filter (0.22 μm) and stored at 4
C). 12. Trypsin-EDTA, in Hank’s Balanced Salt Solution, 0.05% Trypsin, 0.53 mM EDTA, without Ca2+ and Mg2+. 13. 15 mL and 50 mL sterile centrifuge tubes. 14. Culture flasks, 25 cm2, 75 cm2, and 175 cm2. 15. Individually wrapped sterile disposable pipettes. 16. Type 1 UPW. 17. Ethanol, 70%, in spray bottle. 18. Absorbent paper tissues. 19. Waste bottle.

2.3

BBB Modeling

1. All materials in Subheading 2.2. 2. Cell Culture Insert, sterile transparent PET membrane (see Note 2). 3. Sterile 12 and 24 well plates.

2.4 Characterization of the BBB Model

1. Fluorescence spectrometer. 2. Inverted fluorescent microscope. 3. TEER device (Volt/ohm meter) and electrodes. 4. Microplate shaker. 5. Micropipettes, 1–10 μL, 10–100 μL, and 100–1000 μL. 6. Lucifer Yellow (LY) CH dipotassium salt.

In Vitro Human Blood-Brain Barrier Model for Drug Permeability Testing

77

7. Ringer’s HEPES Solution (1) in type 1 UPW: 150 mM NaCl, 3.4 mM CaCl2, 1.2 mM MgCl2, 5.2 mM KCl, 0.5 mM NaHCO3, 2.8 mM glucose, and 10 mM HEPES, pH 7.4. Sterile Ringer’s HEPES by passing through a syringe filter (0.22 μm) and store at 4
C. 8. Tight junction proteins primary antibodies (e.g., Anti-Zonula Occludens-1 (ZO-1), Anti-Claudin 5, Anti-Occludin, etc.) (see Note 3). 9. Secondary antibodies (see Note 4). 10. Ca2+ and Mg2+ free PBS. 11. PBS with Ca2+ and Mg2+ (1): 0.2 g/L KCl, 0.2 g/L KH2PO4, 8 g/L NaCl, 2.16 g/L Na2HPO4.7H2O, 0.2 g/L CaCl2 (anhydrous), 0.98 g/L MgSO4.7H2O in type 1 UPW, pH 7.4. Sterile PBS by passing through a syringe filter (0.22 μm) and store at 4
C. 12. Paraformaldehyde (PFA) solution, 4% in type 1 UPW. 13. NH4Cl solution, 50 mM, in type 1 UPW. 14. Triton X-100 solution, 0.2%, in Ca2+ and Mg2+ free PBS. 15. BSA, 3%, in Triton X-100 solution. 16. Mowiol mounting medium solution (see Note 5). 17. DAPI (40 ,6-Diamidino-2-Phenylindole, Dihydrochloride). 18. Type 1 UPW. 19. Scalpel. 20. Parafilm. 21. Petri dish. 22. Glass slides and coverslips. 23. Ethanol, 70%, in spray bottle. 24. Absorbent paper tissues. 25. Waste bottle. 2.5 Drug Permeability Testing

1. Micropipettes, 1–10 μL, 10–100 μL, and 100–1000 μL. 2. Measurement equipment (e.g., UV-VIS spectroscopy, fluorescent spectroscopy, HPLC, LC/Q-TOF/MS, etc.) suitable for selected drug molecule’s properties, to determine the concentration in abluminal side of the insert. 3. Drug molecule to be tested for BBB permeability. 4. Ca2+ and Mg2+ free PBS. 5. Ringer’s HEPES Solution (1). 6. Ethanol, 70%, in spray bottle. 7. Absorbent paper tissues. 8. Waste bottle.

78

3

Ece Bayir and Aylin Sendemir

Methods Carry out all procedures in Class II Biosafety Cabinet. Swab the hood surface and all the materials by 70% ethanol, and place them immediately in the hood. All liquids to be used on the cells must be preheated to 37
C. All the chemicals to be prepared in the medium must either be added to the medium in the required amount after being sterilized as a powder, or must be added to the medium in the desired amounts after the stock has been prepared in their solvents (e.g., type 1 UPW) in high concentrations and passed through a syringe filter (0.22 μm).

3.1

Cell Culture

1. Prepare the triple co-culture growth medium with DMEM (high glucose) supplemented with 10% (w/w) FBS, 1% (w/w) L-Glutamine, 1% (w/w) sodium pyruvate, 50 μg/mL gentamycin, 18.5 μg/mL putrescine, 5.57 μg/mL insulin, 2 μg/mL heparin, 1.95 μg/mL BSA, 0.5 μg/mL hydrocortisone, 30 ng/mL EGF, 6.3 ng/mL progesterone, 5.2 ng/mL sodium selenite, 5 ng/mL bFGF [17]. Store at 4
C and preheat in 37
C water bath before use. 2. Retrieve the cryotubes of the BMECs, pericytes, and astrocytes from liquid nitrogen, and place them in the 37
C water bath. 3. Transfer the thawed vials in the Class II Biosafety Cabinet, add growth medium dropwise to the thawed cell suspensions, and transfer the suspensions in a 15 mL centrifuge tubes, separately. 4. Centrifuge the cell suspensions at 100  g for 5 min and at 4
C. Discard the supernatant and resuspend the cells in 5 mL fresh growth medium. 5. Transfer the cells to 25 cm2 flasks separately, and put the flasks in the incubator after observing the cells on the inverted microscope. 6. Change the medium every other day until the cells reach approximately 80% confluency. 7. Withdraw the medium and discard it into the waste bottle. Rinse the cells with 2 mL Ca2+ and Mg2+ free PBS twice. 8. Add 1 mL Trypsin-EDTA solution and return the cells in the incubator for 1 min. 9. Observe the cells on the inverted microscope. Dilute the trypsin-EDTA with 4 mL fresh growth media or FBS after the cells detach from the surface. 10. Transfer the cells in a centrifuge tube, and centrifuge them for 5 min at 100  g and at 4
C. 11. Discard the supernatant and resuspend the cells in 15 mL fresh growth medium.

In Vitro Human Blood-Brain Barrier Model for Drug Permeability Testing

79

12. Transfer the cells to 75 cm2 flasks and incubate after observing the cells by the inverted microscope. 13. Subculture the cells until they reach the desired number (depending on the number of experimental groups). 14. Count the cells by the hemocytometer or electronic cell counter after detaching cells. 15. Resuspend the cells at desired concentrations (3  105 BMECs/cm2, and BMECs:Astrocytes:Pericytes ratio to be: 1:3:5) in a desired volume of growth medium. 3.2

BBB Modeling

1. Condition the cell culture inserts with growth medium for at least 2 h. 2. Turn the cell culture inserts upside down in a 12-well plate, and fill the plate with co-culture growth medium to the level of the inserts (Fig. 1). 3. Seed the pericytes onto the underside of the insert membrane (Fig. 1) in 50 μL volume of growth medium and place the 12-well plate in the incubator for least 4 h (see Note 6). 4. Seed the astrocytes onto the pericytes (Fig. 1) in 50 μL volume of growth medium and incubate the plate overnight (see Note 6). 5. Turn the cell culture inserts and transfer them in a 24-well plate. 6. Seed the BMECs in 100 μL volume of growth medium on upside of the cell culture inserts and return the plate to the incubator (Fig. 1). 7. Continue co-culture for up to 5 days.

3.3 Characterization of the BBB Model

1. Place the TEER electrodes in 70% ethanol for 15 min for disinfection.

3.3.1 TEER Measurement

2. Put the electrodes in growth medium in the Class II Biosafety Cabinet after disinfection and wait until the TEER device is calibrated to 0 Ω. 3. Measure the TEER values of the blank media by inserting the electrodes into the cell culture inserts that only contain growth media for background measurements (see Note 7). 4. Wash the electrodes in clean medium after each measurement. 5. Measure the TEER values of triple co-culture inserts (at least 3 repetitions for each experimental group). It is accepted that in vitro models with a TEER value of >150 Ω.cm2 are suitable for pharmaceutical studies [18].

80

Ece Bayir and Aylin Sendemir

Fig. 1 Cell seeding steps on cell culture inserts. (a) Pericytes are seeded on the abluminal side of the BBB model first. (b) Astrocytes are seeded on the attached pericytes after at least 4 h. (c) The inserts are transferred to 24-well plates, and endothelial cells are seeded on the luminal side of the model after the cells are attached (overnight). (d) The final BBB model 3.3.2 Lucifer Yellow (LY) Permeability

1. Prepare 0.1 M LY solution in Ringer’s HEPES and pass through a syringe filter (0.22 μm) for sterilization. Store at room temperature and protect from light. 2. Dilute 0.1 M LY solution to at least 5 different concentrations between 1  104 and 1  107 M, and measure at 428–540 nm (excitation-emission) on fluorescence spectrometer. Draw a calibration curve with the fluorescence values corresponding to the LY concentrations. 3. Withdraw the growth medium in the luminal and abluminal sides of the cell culture inserts and discard in waste bottle in the Class II Biosafety Cabinet. 4. Rinse the cells by prewarmed sterile PBS twice. Add 500 mL of Ringer’s HEPES solution to the abluminal side of the inserts. 5. Add 200 mL of 0.1 M LY solution into the luminal side of the inserts (at least 3 repetitions for each experimental group) and incubate the 24-well plate for a total of 120 min. After 30 min, take desired volume of the Ringer’s HEPES solution from the abluminal side of the inserts for reading. Return the plate to the incubator and repeat the measurement after 60 min and 120 min after adding LY solution. Read samples of the Ringer’s HEPES solution from each group at 428–540 nm (excitationemission) on fluorescent spectroscope.

In Vitro Human Blood-Brain Barrier Model for Drug Permeability Testing

81

6. Calculate permeability values of LY from in vitro BBB model (see Note 8). The model can be used for drug permeability testing if LY molecules have 150 Ω.cm2, permeability value of LY is 97%) primary brain endothelial cells (BECs) is produced by plating 5  105 cells/cm2 in the upper chamber of a 24-well tissue culture inserts (BD BioCoat, Catalog # 356408) pre-coated with rat tail collagen I and fibronectin. BECs are >98% pure, as measured by staining for the endothelial cell specific markers von Willebrand Factor (vWF) and CD105, and negative immunostaining for the glial fibrillar acidic protein (astrocytes), CD11b (macrophage/microglia), smooth muscle cell α-actin (SMA), and platelet-derived growth factor receptor β and CD13 (pericytes). Early passage P2-P5 cells and TER value of >200 Ω cm2 are generally recommended. 2. Endothelial complete medium: BECs are maintained in endothelial medium with supplements (ScienCell, complete kit, Catalog #1001). 3. Isothiocyanate (FITC) labeled dextran (e.g., 40 kDa; Invitrogen, Catalog D1845). 4. 96-well microplates for fluorescence-based assays (Invitrogen, Catalog M33089). 5. Phosphate-buffered saline (PBS), for cell culture, without calcium and magnesium (Invitrogen, Catalog 10010023).

2.2 Materials for BBB Permeability and Transport Measurement In Vivo

Buffers. Mammalian Ringer’s solution is used as the solvent for making the solution with the fluorescently labeled solutes (perfusate). All the solutions additionally contain 10 mg/mL bovine serum albumin (BSA) or 1%BSA to simulate the normal oncotic pressure in the blood. The solutions are made at the room temperature 25  C and warmed up to 37  C before perfusion into the rat brain through the carotid artery. The following quantity of the solution for each solute is made for the control experiments using one to two rats. Artificial cerebrospinal fluid (ACSF) is used to superfuse the surface of the rat scalp to remove the heat generated during the grinding. 1. Mammalian Ringer solution: Dissolve 7.714 g NaCl, 0.3432 KCl, 0.294 CaCl22H2O, 0.2957 g MgSO47H2O, 0.99 g glucose, 0.42 g NaHCO3, 2.37 g Hepes (salt), and 2.6 g Hepes (acid) in 1 L double-distilled water (13). Store at 4  C for a week. 2. 1% BSA solution: Dissolve 500 mg BSA in 50 mL Mammalian Ringer to make 50 mL 10 mg/mL BSA (1% BSA) solution. The solution is buffered to pH 7.4–7.5 [16]. 3. Fluorescently labeled solute solutions: FITC or TRITC or other fluorescently labeled solutes, e.g., dextrans with various

108

Bingmei M. Fu et al.

molecular weights, sodium fluorescein, TAMRA, are often used to represent solutes with a range of sizes. We use sodium fluorescein (NaFl, MW 376) and FITC-dextran 70k as a typical small and a large-sized solute to demonstrate the permeability measurement method. For sodium fluorescein and FITClabeled solutes, the excitation/emission wavelengths are set to 485/520 nm in the generic fluorescence microscopy (13) and 820 nm for the excitation in the multiphoton microscopy [12–15]. 4. 50 mL sodium fluorescein (NaFl) at 0.1 mg/mL: Dissolve 5 mg NaFl in 50 mL 1%BSA solution, and filter the solution through a 0.2 μm syringe filter (0.1 mg/mL sodium fluorescein is in the linear range of the concentration vs. intensity curve for our experimental settings). 5. 50 mL FITC-dextran 70k at 1 mg/mL: Dissolve 50 mg dextran 70k in into 50 mL 1% BSA and filter the solution through a 0.2 μm syringe filter (1 mg/mL FITC-dextran is in the linear range of the concentration vs. intensity curve for our experimental settings). 6. ACSF solution: Dissolve 6.4576 g NaCl, 0.3507 KCl, 0.3675 CaCl22H2O, 0.308 g MgSO47H2O, 2.1 g NaHCO3, 0.1497 g KH2PO4, 1.7775 g Hepes (salt), and 1.95 g Hepes (acid) in 1 L double-distilled water [11]. The solution is buffered to 7.4  0.5. Store at 4  C for a week.

3

Methods Part I: BBB Permeability Measurement In Vitro: Carry out all procedures in biosafety hood at room temperature, unless otherwise specified (see Note 1). Prepare all solutions using ultrapure water and analytical grade reagents. Prepare fresh work solutions, and follow all disposal regulations when disposing waste.

3.1 Apply Experimental Treatments

Starvation with low serum (endothelial basal medium with 0.1–0.5% serum), or addition of cyclic adenosine monophosphate (cAMP) [23] or retinoic acid [24], or coculture with astrocytes [25] or pericytes [26, 27], has been shown to reliably increase the barrier functions of in vitro BBB. Either replace the growth medium in the bottom chamber or transfer the insert to a receiver well with pre-seeded coculture cells, and then cover the plate and incubate the cells (see Note 2).

BBB Permeability and Transport in Brain

3.2 Dextran Permeability Measurement

109

1. Prepare FITC-dextran in growth medium to a final concentration of 0.1 mg/mL. 200 μL of volume is needed for each insert. 2. Prepare a 96-well fluorescence-based microplate, and add 100 μL PBS in 8 wells for every insert need to be tested (including a cell-free filter insert). If necessary, label the wells in two sets of four wells, one for samples collected from upper chamber at different time points and one for samples from lower chamber. 3. Add 5 μL of FITC-dextran medium from Subheading 3.2 step 1 to the 96-well fluorescence microplate as first time point of upper chamber for each insert need to be tested. 4. Transfer the plate with inserts to the biosafety hood, and add 5 μL of the medium from the bottom chamber of each inserts (including a cell-free filter insert) to the 96-well fluorescence microplate as first time point of bottom chamber. Carefully remove the medium from the insert, without the pipette tip touching the membrane or drying the membrane. 5. Add 200 μL of FITC-dextran medium to the insert gently, without the pipette tip touching the membrane. Move the plate back to the incubator. Start the timer. 6. Every 10 min collect 5 μL of medium from both the upper chamber inside the insert and the bottom chamber, and transfer to the designated wells in 96-well fluorescence microplate. Collect for the first 30 min (4 time points total including first time point collected in Subheadings 3.2, steps 3 and 4). 7. Keep the inserts in PBS for phase-contrast imaging or immunofluorescent staining of vascular and tight junction markers if needed.

3.3 Calculating the Permeability of the Endothelial Barrier

The BBB permeability to dextran is expressed as a permeability coefficient in cm/s [27]. 1. Read the 96-well fluorescence microplate using a plate reader with filter setting at 488 nm for excitation and 525 nm for emission or compatible settings recommended by the manufacturer (see Note 3). 2. For each time point, calculate the ratio of fluorescence between upper chamber and lower chamber (Fupper/Flower). 3. The volume cleared (ΔVc) of each time point is calculated using Eq. 1 [26, 27]: ΔV c ¼ C lower  V lower =C upper where Cupper and Clower are FITC-labeled dextran concentrations in upper and lower chambers, respectively, Vlower is the

110

Bingmei M. Fu et al.

volume in lower chamber (~500 μL), and Cupper/Clower is equivalent to the Fupper/Flower measured from fluorescence reading in Subheading 3.3, step 2. 4. Plot the ΔVc against time (0–30 min) over time in Excel, and the permeability surface area (PS) product can be obtained from the slope by linear regression. The permeability coefficient (P) of the insert (cell and filter) can then be calculated by Eq. 2 [26, 27]: P cellþfilter ¼ PS=s where s is the surface area of the filter (0.3 cm2 for 24-well inserts). 5. The permeability coefficient of cells within the insert (Pcell) can be obtained by correcting the overall permeability coefficient (Pcell + filter) for that of the cell-free filter (Pfilter) using Eq. 3 [26, 27] (see Note 4): 1=P cell ¼ 1=P cellþfilter  1=P filter where Pfilter is determined using the cell-free filter insert only. Part II: BBB Permeability Measurement In Vivo: The method for quantification of the solute permeability (P) is from [13, 15, 16], and that for quantification of the effective brain tissue diffusion coefficient (Deff) is from [14]. 3.4 Calibration of Optical Settings of Microscope for Linear Range of Concentration vs. Intensity of Fluorescence Probe Solutions

The premise for using the fluorescence intensity as an indicator of the solute concentration is that the intensity changes linearly with the concentration measured by the instrument (e.g., intravital microscope) for the range of concentrations in the experiment. We use here either a Nikon TE2000-E inverted fluorescence microscope (objective lens 20x/NA0.75, spatial resolution ~0.5 μm/ pixel and temporal resolution three frame/s for image size 1024  1376) with a 12-bit CCD camera (Sensicam QE, Cooke, MI) for measuring the solute permeability P of pial microvessels or an Ultima multiphoton microscopy system (Prairie Tech. Inc. WI) with an objective lens 40x/NA0.8 (spatial resolution ~0.5 μm/ pixel and temporal resolution ~1 frame/s for image size 512  512) for measuring P of cerebral microvessels 100–200 μm below pia mater. 1. Make sodium fluorescein (NaFl) solution 0.5 mL each at 0.2, 0.1, 0.05, 0.025, and 0.0125 mg/mL, respectively, or FITCdextran 70k 0.5 mL each at 2, 1, 0.5, 0.25, and 0.125 mg/mL, respectively.

BBB Permeability and Transport in Brain

111

2. Make a chamber with a gap of 170 μm by using four 170 μm thick glass cover slips. Two large ones (24  50 mm) for the top and bottom and two small ones (22  22 mm) sandwiched between the large ones, ~1 cm apart (see Note 5). Solutions of fluorescence probe (NaFl or FITC-dextran-70k) are applied to fill the gap by capillarity. 3. The chamber filled with the fluorescence probe solution is put under the microscope and the intensity is measured. The measuring window is ~250  300 μm for the Nikon microscope and ~250  250 μm for the multiphoton microscope, similar size to that in the in vivo experiment. Adjust the optical settings of the microscope to be sure that the measured intensity of the fluorescence probe solution is linearly correlated with its concentration in the range shown in step 1. This setting and the concentration in the linear range will be used in the in vivo experiment (see Note 6). 3.5 Determination of the Depth of Light Collection for the Imaging System

Since the measurements of fluorescence intensity for cerebral microvessels are performed on thick tissue, collection of the light from out of focus region would contribute to the measured value of intensity. To determine the depth of light collection of our system, in vitro experiments similar to [28] are performed but using an imaging system to determine the fluorescence intensity. 1. Four chambers with different gap depths, 50 μm from Intracellular Imaging Inc. (Cincinnati, OH), 100 μm (hemocytometer), 170 μm chamber formed by cover slips (described in the previous Subheading 3.1), and 340 μm chamber with two cover slips stacked, are used to measure fluorescence intensity of sodium fluorescein solutions, which are filled in the chambers by capillarity. 2. The experiment instrument settings are the same as in in vitro linear calibration experiments as well as in in vivo permeability measurement (objective lens 20x/NA ¼ 0.75 for the Nikon TE2000-E microscope and 40x/NA0.8 for the multiphoton microscope). With the concentration-depth product kept constant, the solutions of fluorescent probes are diluted according to the different depth of the chambers [28]. The intensity is measured for a window area ~250  300 μm for the Nikon microscope and ~250  250 μm for the multiphoton microscope, and the focus is on the top surface of the solution. Results are shown in Fig. 1. Intensities for 50 and 100 μm deep chambers remain almost the same. However, for the chambers with depths larger than 100 μm, i.e., 170 and 340 μm chambers, the intensity decreased with increase of the depth of the chambers. The reason is that our experiment system can only collect the light from the solution in a smaller

112

Bingmei M. Fu et al.

Fig. 1 Depth of light collection. Samples of sodium fluorescein solution are prepared such that concentration-depth product is maintained constant. The total fluorescence intensity, which is normalized by the value for the 50 μm depth chamber, is plotted for the chambers with the depth 50, 100, 170, and 340 μm. The continuous line is the curve fitting for the total intensity using a light collection index function in Yuan et al. [16]. (Reprint with the permission from ref. [16], Copyright 2009, Elsevier Inc.)

thickness region for the 170 and 340 μm chambers. Using a light collection index function defined in [28], the curve fitting is done for the measured total intensity in the chambers of different depths. We find that for the Nikon microscope system with objective lens 20x/NA0.75, the depth of light collection is 95 μm (z0 ¼ 95 μm in their index function), with this depth the imaging system can collect all the light. Beyond this depth, the system can collect the light either partially or none. Similarly, the depth of light collection is 67 μm for our multiphoton microscope with objective lens 40x/NA0.8 (see Note 7). 3.6 Animal Preparation

1. Adult Sprague-Dawley rats (250–300 g, 3–4 months) are used. 2. After anesthesia, the rat is kept warm on a heating pad. The skull in the region of interest is exposed by shaving off the hair and cutting away the skin and connective tissue. A section of left or right frontoparietal bone, approximately ~5 mm ~5 mm, is carefully ground with a high-speed micro-grinder (0–50,000 rpm, DLT 50KBU; Brasseler USA, GA) until a part of it (~2 mm  ~2 mm) becomes soft and translucent (~100 μm thickness). During the process, artificial cerebrospinal fluid (ACSF) with the room temperature is applied to the surface of the skull to remove the heat due to

BBB Permeability and Transport in Brain

113

grinding. After grinding, the right or left carotid artery (on the same side of the thinned scalp) is cannulated with PE50 tubing. 3. If use the Nikon TE2000-E inverted fluorescence microscope, after grinding and cannulating, the rat is laid face up on a tray with its thinned section of the head placed in a shallow chamber formed by a glass cover slip. The thinned section is observed under the microscope, and the pial microvessels are focused by carefully adjusting the focusing knob. If use the upright Ultima multiphoton microscopy system, the rat is placed on a stereotaxic alignment system (SAS 597; David Kopf Instruments, Tujunga, CA), and its head is fixed with two ear bars and a mouth clamp. The thinned part of the scalp is placed underneath the water immerging objective lens. 3.7 Image Acquisition 3.7.1 2-D Image Collection for the Measurement of Solute Permeability of the BBB

The microvessels (post-capillary venules of diameter 20–40 μm, see Note 8) are observed by the above microscope system through the thinned part of the rat scalp; the fluorescent solution is introduced into the cerebral circulation via the left or right carotid artery (on the same side of the thinned part) by a syringe pump at a constant rate of 3 mL/min (see Note 9), while the images of the fluorescent solution-filled microvessels and nearby brain tissue are simultaneously collected by the imaging systems. The Nikon system collects the images (~512  688 μm, ~0.5 μm/pixel and ~3 frame/s) of the pial microvessels, while the multiphoton system collects the images (~250  250 μm, ~0.5 μm/pixel and ~1 frame/ s) of the cerebral microvessels ~100–200 μm below the pia mater (Fig. 2).

3.7.2 3-D Image Collection for Determination of Solute Permeability and Effective Diffusion Coefficient in Brain Tissue

The 3-D images are collected by the multiphoton microscope for a region of interest (ROI) with a volume of ~200 μm  8 μm  100 μm (x, y, z) (Fig. 3) at a rate of 5–15 s per image (see Note 10). The corresponding pixel sizes were ~428 pixel  17 pixel  100 pixels, resulting a spatial resolution of ~0.47 μm  0.47 μm  1 μm in x, y, and z directions. The collected images of the ROI are then transferred to an image acquisition and analysis workstation for determining the P and Deff (Fig. 4).

3.8

The images are analyzed using ImageJ (National Institutes of Health).

Image Analysis

1. For the 2-D images, the analysis is straightforward, for the intensity of a ROI as a function of time (Fig. 2). 2. For the 3-D images, the collected images are first reconstructed into a segment of 200 μm  100 μm cross-sectional area (x-z) with 8 μm thickness. The temporal and spatial solute intensity (concentration) profiles I (t, xt, zt) surrounding a microvessel in this cross-sectional area with 8 μm thickness of the brain tissue are determined by the ImageJ program.

114

Bingmei M. Fu et al.

Fig. 2 Determination of the BBB solute permeability P from 2-D images. (a) Illustration of a 2-D image comprising several microvessels. The measuring window (yellow framed area) is drawn on top of a chosen microvessel for determining the P of that vessel. (b) Total fluorescence intensity in the measuring window as a function of the perfusion time. Fluorescence intensity in the figure is proportional to the total amount of the solutes accumulated in the measuring region surrounding the microvessel. The slope of the regression line over the initial linear accumulation (dI/dt)0 (red line) is used to determine the solute permeability P ¼ 1/Δ I0  (dI/dt)0  r/2, where ΔI0 (black line with arrowheads) is the step intensity increase when the dye just fills up the vessel lumen and r is the radius of the vessel (Reprint with the permission from ref. [15], Copyright 2020, Springer Link) 3.9 Determination of Solute Permeability of BBB (PBBB) from Collected Images

Figure 2 demonstrates how to determine PBBB from the 2-D images collected for the ROI comprising several microvessels (either the pial microvessels at the brain surface or the cerebral microvessels ~100–200 μm below the pia mater).

3.9.1 Determination of Solute Permeability of BBB (PBBB) from 2-D Images

1. Find a microvessel in the image, and draw a measuring window about 30–60 μm wide and 50–100 μm long (orange frame in Fig. 2a) (see Note 11). 2. Plot the increasing intensity of the fluorescent probe in the measuring window as a function of perfusion time (Fig. 2b). The slope of the regression line over the initial linear accumulation (dI/dt)0 (red line) is used to determine the solute permeability P ¼ 1/ΔI0  (dI/dt)0  r/2, where ΔI0 (black line with arrowheads) is the step intensity increase when the dye just fills up the vessel lumen and r is the radius of the vessel.

3.9.2 Determination of Solute Permeability of BBB (PBBB) from 3-D Images

1. Figure 4a shows a cross-sectional image (x-z) of a rat cerebral microvessel filled with a solution of fluorescently labeled solutes and the surrounding brain tissue. The orange dashed line circled region is the ROI to determine P (see Note 12).

BBB Permeability and Transport in Brain

115

Fig. 3 Illustration of the scanning region and the orientation of a microvessel in the brain tissue using multiphoton microscopy to determine the BBB solute permeability and its brain tissue diffusion coefficient. x-z is the cross-sectional plane of the microvessel, and y is the axial direction of the vessel. The ROI (scanning region) has a size of ~200 μm x ~ 8 μm x ~ 100 μm (x, y, z) (Reprint with the permission from ref. [14], Copyright 2014, American Society of Mechanical Engineers)

2. The total intensity in the ROI as a function of perfusion time. The slope of the regression line over the initial linear accumulation is used for the estimation of solute permeability P ¼ 1/ I0  (dI/dt)0  r/2. I0 is fluorescence intensity in the vessel lumen while r is the radius of the vessel (Fig. 4b). The determined P is used in the boundary condition (in Eq. 4) for solving the unsteady transport equation to predict the spatialtemporal solute tissue distribution profiles surrounding a microvessel (see Note 13). 3.10 Determination of Effective Solute Diffusion Coefficient in Brain Tissue (Deff) from 3-D Images

Figure 4a, c illustrates how to determine Deff from the collected images 1. Eight straight white lines in different radial directions are drawn from the center of the vessel (Fig. 4a). 2. The averaged fluorescence intensity along the eight lines at different times (t ¼ 25, 50, and 75 s) is plotted from the vessel wall (Fig. 4c). The smooth lines are the best fitting curves of the model prediction at the corresponding times when the

116

Bingmei M. Fu et al.

Fig. 4 Determination of BBB solute permeability P and effective solute diffusion coefficient Deff in rat brain tissue from 3-D images. (a) The cross-sectional image of a rat cerebral microvessel filled with fluorescent probes and its surrounding brain tissue. The dashed orange line enclosed circle is the ROI for determining

BBB Permeability and Transport in Brain

117

proper Deff is chosen (see Note 13). Deff/Dfree ¼ 0.15 (Dfree is the solute diffusion coefficient in an aqueous solution at 37  C) is the best fitting value for this run of the experiment. 3.11 Corrections for Influence of Red Blood Cells, Free Dye, and Solvent Drag on BBB Permeability

The above determined BBB solute permeability is the uncorrected permeability. Three factors, from red blood cells, free dye, and solvent drag, need to be corrected to find the true diffusive permeability defined in Kedem-Katchalsky equations (see introduction). 1. The fluorescence dye solution is injected into the brain at the rate of 3 ml/min, the same as the normal blood perfusion rate at the carotid artery. Although at this perfusion rate the blood is assumed to be replaced by the fluorescence solution, there is still residue blood (red blood cells, RBCs) in the cerebral microvessels, which would overestimate the measured BBB permeability by ~11%, as estimated in [12]. 2. In addition to RBCs, free dye would overestimate the permeability to fluorescently labeled solutes. The influence of the free dye on the solute permeability is estimated by using equation Pcorrect ¼ [1/(1F)] Pmeasure – [F/(1F)] Pfreedye, where Pmeasure was the measured permeability and Pfreedye is similar to PNaFl since the molecular weights of FITC (389.4) is close to that of sodium fluorescein (NaFl, 376). F is the intensity fraction of the free dye to the fluorescently labeled probe in the solution. F is less than 1/1000 in the commonly used FITCconjugated solute solutions [12–16]. Pcorrect is the corrected apparent permeability P. 3. The above apparent permeability P corrected for the RBCs and free dye still overestimates the true diffusive solute permeability Pd due to the coupling of solute flux with water flow (solvent drag). The Pd is calculated by using the following equations: P ¼ Pd

Pe þ L p ð1  σ ÞΔpeff exp ðP e  1Þ

ä Fig. 4 (continued) vessel permeability P. Eight straight white lines in different radial directions are drawn from the center of the vessel to determine Deff. (b) The total intensity in the ROI as a function of perfusion time. The slope of the regression line over the initial linear accumulation is used for the estimation of solute permeability P ¼ 1/I0  (dI/dt)0  r/2. I0 is fluorescence intensity in the vessel lumen, while r is the radius of the vessel. (c) The averaged fluorescence intensity along the eight lines at different times (t ¼ 25, 50 and 75 s) are plotted from the vessel wall. The smooth lines are the best fitting curves of the model prediction at the corresponding times when the proper Deff is chosen. Deff/Dfree ¼ 0.15 (Dfree is the solute diffusion coefficient in an aqueous solution at 37  C) is the best fitting value for this run of the experiment. (Reprint with the permission from ref. [14], Copyright 2014, American Society of Mechanical Engineers)

118

Bingmei M. Fu et al.

L p ð1  σ ÞΔpeff Pd where P is the corrected apparent permeability, Pe is the Peclet number, Lp is the hydraulic conductivity of the microvessel, which is ~2.0  109 cm/s/cm H2O for the cerebral microvessels [29], σ is the reflection coefficient of the microvessel to the solute, and Δpeff is the effective filtration pressure across the microvessel wall, obtained from: Pe ¼

Δpeff ¼ Δp  σ albumin Δπ albumin  σ dyesolute Δπ dyesolute where Δp and Δπ are the hydrostatic and oncotic pressure differences across the microvessel wall. The superscript dye-solute is NaFl or Dex-70k, in our examples. σ of rat cerebral microvessels to the test solutes is estimated based on previous studies [12] according to the molecule sizes. σ dextran70k (the same as σ albumin) and σ NaFl were estimated to be 0.95 and 0.1, respectively. Δp in the cerebral microvessel was ~10 cm H2O; Δπ albumin was 3.6 cm H2O for 10 mg/mL BSA [12].

4

Notes 1. All procedures should be performed in biosafety hood with laminar flow, and handling of the inserts should be carefully done with autoclaved forceps to avoid potential contaminations. 2. The insert with cells should be handled with autoclaved forceps. 3. Although all the perimeters including the dextran dilution have been previously tested, it may still be worthwhile running a standard curve for fluorescent reading of dextran at different dilutions to make sure that the concentration (C) and fluorescence reading (F) from the plate reader will be in a linear range. 4. The permeability coefficient also depends on the size of a molecular, or its molecule weight [30]; therefore using different sizes of dextran will result in differences in permeability coefficient results [27]. 5. The chamber is cleaned by 70% ethanol before applying the solution. 6. For example, sodium fluorescein of 0.1 mg/mL, in the linear range of 0.0125–0.2 mg/mL, and FITC-dextran 70k of 1 mg/ mL, in the linear range of 0.125–2 mg/mL, will be used in the in vivo experiment.

BBB Permeability and Transport in Brain

119

7. Within this depth of the light collection, the fluorescence intensity is proportional to the total number of fluorescent molecules and is independent of the chamber depth. This narrow depth of light collection in our system allows us to minimize the influence from the light coming from other parts of the brain tissue, especially from the meninges for the pial microvessels. It is very important to use the objective lens with proper magnification and numerical aperture to achieve narrow but enough depth of light collection to collect correct samples with minimum light contamination from irrelevant parts. 8. The post-capillary venule is most permeable among all microvessels. We follow the method described in [16] for identifying post-capillary venules. 9. About 3 mL/min is the mean blood flow rate at the carotid artery for this sized rat. 10. The collecting time is related to the image quality. 5–15 s per ROI in our experiment is good enough for the image quality for determining the BBB solute permeability and effective diffusion coefficient in brain tissue. 11. The size and placement of the measuring window are chosen to satisfy that (a) the vessel segment is straight, (b) the dye does not spread out of the window during the time for P measurement (5–30 s for the size of dyes used in our experiment), and (c) no contamination of the dye from the neighboring vessels. 12. The circumference of the orange dashed line enclosed ROI is chosen as 10–30 μm from the vessel perimeter to avoid contamination from the adjacent vessels but large enough to include the spreading dye from the vessel lumen during the period for the P measurement. 13. Deff is determined by fitting the temporal and spatial intensity curves by an unsteady mathematical model for solute transport in the tissue space:  2  ∂C t ∂ C t 1 ∂C t ∂C t þ ¼ D eff  χu r ∂r ∂t ∂r 2 ∂r

ð1Þ

where Ct (t, r) is the concentration of solutes in the tissue space, Deff is the effective diffusion coefficient of solutes in tissue, and r is the distance from the vessel center. χ is the retardation coefficient of a solute in the tissue, estimated as 0.1–1 for solutes under study [16]. u is the interstitial fluid velocity in brain tissue. The Peclet number Pet in the tissue is:

120

Bingmei M. Fu et al.

χVL t ð2Þ D eff Here Lt is the characteristic length for the solute tissue transport, which is the mean half distance (~20 μm) between adjacent microvessels [14]; V is the characteristic interstitial fluid velocity, which is approximated by the outflow velocity from the vessel wall LpΔpeff. Lp is the hydraulic conductivity of the microvessel, ~2  109 cm/s/cm H2O [29], while Δpeff is the effective pressure difference across the vessel wall, which is less than 10 cm H2O [16]. For the size range of solutes in this study, Deff is estimated in the range of 106 to 108 cm2/s; Pet was calculated as in the order of 105 to 103. Thus, the convection part can be neglected in Eq. 1. Equation 1 becomes: P et ¼

 2  ∂C t ∂ C t 1 ∂C t þ ¼ D eff r ∂r ∂t ∂r 2

ð3Þ

The boundary conditions for Eq. 3 are at the vessel wall r ¼ a ∂C t ∂r midway between adjacent vessels r ¼ b P ðC lumen  C t Þ ¼ D eff

∂C t ¼0 ∂r

ð4Þ

ð5Þ

The initial condition is: ð6Þ t ¼ 0, C t ð0, r Þ ¼ 0 where Clumen is the solute concentration in the vessel lumen and P is the microvessel solute permeability, both of which can be determined from the collected images. The only unknown parameter in Eqs. 3–6 is Deff. Solving above Eq. 3 with an assumed value of Deff by Matlab, we obtained the theoretical solute tissue concentration profiles Ct(t, rt) (rt ¼ 0 is at the vessel wall). The Deff is determined by the best curve fitting of the model predictions (smooth lines in Fig. 4c) to the measured profiles (crooked lines in Fig. 4c).

BBB Permeability and Transport in Brain

121

Acknowledgments The author would like to thank the funding support from the National Institutes of Health R01NS101362 (B. Fu) and R01AG064798 (D. Zhu). References 1. Abbott NJ, Patabendige AA, Dolman DE, Yusof SR, Begley DJ (2010) Structure and function of the blood-brain barrier. Neurobiol Dis 37(1):13–25. https://doi.org/10.1016/j. nbd.2009.07.030 2. Farkas E, Luiten PG (2001) Cerebral microvascular pathology in aging and Alzheimer’s disease. Prog Neurobiol 64(6):575–611. https:// doi.org/10.1016/s0301-0082(00)00068-x 3. Nicolazzo JA, Charman SA, Charman WN (2006) Methods to assess drug permeability across the blood-brain barrier. J Pharm Pharmacol 58(3):281–293. https://doi.org/10. 1211/jpp.58.3.0001 4. Fu BM (2018) Transport across the bloodbrain barrier. Adv Exp Med Biol 1097:235–259. https://doi.org/10.1007/ 978-3-319-96445-4_13 5. Cornford EM, Young D, Paxton JW, Sofia RD (1992) Blood-brain barrier penetration of felbamate. Epilepsia 33(5):944–954. https://doi. org/10.1111/j.1528-1157.1992.tb02205.x 6. Zlokovic BV, Begley DJ, Djuricic BM, Mitrovic DM (1986) Measurement of solute transport across the blood-brain barrier in the perfused Guinea pig brain: method and application to N-methyl-alpha-aminoisobutyric acid. J Neurochem 46(5):1444–1451. https://doi.org/ 10.1111/j.1471-4159.1986.tb01760.x 7. de Lange EC, de Boer BA, Breimer DD (1999) Microdialysis for pharmacokinetic analysis of drug transport to the brain. Adv Drug Deliv Rev 36(2–3):211–227. https://doi.org/10. 1016/s0169-409x(98)00089-1 8. Elsinga PH, Hendrikse NH, Bart J, Vaalburg W, van Waarde A (2004) PET studies on P-glycoprotein function in the blood-brain barrier: how it affects uptake and binding of drugs within the CNS. Curr Pharm Des 10 (13):1493–1503. https://doi.org/10.2174/ 1381612043384736 9. Wang R, Ashwal S, Tone B, Tian HR, Badaut J, Rasmussen A, Obenaus A (2007) Albumin reduces blood-brain barrier permeability but does not alter infarct size in a rat model of neonatal stroke. Pediatr Res 62(3):261–266. https://doi.org/10.1203/PDR. 0b013e318123f757

10. Gaber MW, Yuan H, Killmar JT, Naimark MD, Kiani MF, Merchant TE (2004) An intravital microscopy study of radiation-induced changes in permeability and leukocyte-endothelial cell interactions in the microvessels of the rat pia mater and cremaster muscle. Brain Res Brain Res Protoc 13(1):1–10. https://doi.org/10. 1016/j.brainresprot.2003.11.005 11. Easton AS, Fraser PA (1994) Variable restriction of albumin diffusion across inflamed cerebral microvessels of the anaesthetized rat. J Physiol 475(1):147–157. https://doi.org/10. 1113/jphysiol.1994.sp020056 12. Shi L, Palacio-Mancheno P, Badami J, Shin DW, Zeng M, Cardoso L, Tu R, Fu BM (2014) Quantification of transient increase of the blood-brain barrier permeability to macromolecules by optimized focused ultrasound combined with microbubbles. Int J Nanomedicine 9:4437–4448. https://doi.org/10. 2147/IJN.S68882 13. Shi L, Zeng M, Fu BM (2014) Temporal effects of vascular endothelial growth factor and 3,5-cyclic monophosphate on bloodbrain barrier solute permeability in vivo. J Neurosci Res 92(12):1678–1689. https://doi. org/10.1002/jnr.23457 14. Shi L, Zeng M, Sun Y, Fu BM (2014) Quantification of blood-brain barrier solute permeability and brain transport by multiphoton microscopy. J Biomech Eng 136(3):031005. https://doi.org/10.1115/1.4025892 15. Shin DW, Fan J, Luu E, Khalid W, Xia Y, Khadka N, Bikson M, Fu BM (2020) In vivo modulation of the blood-brain barrier permeability by transcranial direct current stimulation (tDCS). Ann Biomed Eng 48(4):1256–1270. https://doi.org/10.1007/s10439-02002447-7 16. Yuan W, Lv Y, Zeng M, Fu BM (2009) Non-invasive measurement of solute permeability in cerebral microvessels of the rat. Microvasc Res 77(2):166–173. https://doi. org/10.1016/j.mvr.2008.08.004 17. Crone C, Olesen SP (1982) Electrical resistance of brain microvascular endothelium. Brain Res 241(1):49–55. https://doi.org/10. 1016/0006-8993(82)91227-6

122

Bingmei M. Fu et al.

18. Pardridge WM (1998) CNS drug design based on principles of blood-brain barrier transport. J Neurochem 70(5):1781–1792. https://doi. org/10.1046/j.1471-4159.1998. 70051781.x 19. Patlak CS, Fenstermacher JD (1975) Measurements of dog blood-brain transfer constants by ventriculocisternal perfusion. Am J Phys 229 (4):877–884. https://doi.org/10.1152/ ajplegacy.1975.229.4.877 20. Nicholson C, Phillips JM (1981) Ion diffusion modified by tortuosity and volume fraction in the extracellular microenvironment of the rat cerebellum. J Physiol 321:225–257. https:// doi.org/10.1113/jphysiol.1981.sp013981 21. Thorne RG, Nicholson C (2006) In vivo diffusion analysis with quantum dots and dextrans predicts the width of brain extracellular space. Proc Natl Acad Sci U S A 103(14):5567–5572. https://doi.org/10.1073/pnas.0509425103 22. Stroh M, Zipfel WR, Williams RM, Webb WW, Saltzman WM (2003) Diffusion of nerve growth factor in rat striatum as determined by multiphoton microscopy. Biophys J 85 (1):581–588. https://doi.org/10.1016/ S0006-3495(03)74502-0 23. Ishizaki T, Chiba H, Kojima T, Fujibe M, Soma T, Miyajima H, Nagasawa K, Wada I, Sawada N (2003) Cyclic AMP induces phosphorylation of claudin-5 immunoprecipitates and expression of claudin-5 gene in bloodbrain-barrier endothelial cells via protein kinase A-dependent and -independent pathways. Exp Cell Res 290(2):275–288. https://doi.org/ 10.1016/s0014-4827(03)00354-9 24. Lippmann ES, Al-Ahmad A, Azarin SM, Palecek SP, Shusta EV (2014) A retinoic acidenhanced, multicellular human blood-brain barrier model derived from stem cell sources. Sci Rep 4:4160. https://doi.org/10.1038/ srep04160

25. Ma SH, Lepak LA, Hussain RJ, Shain W, Shuler ML (2005) An endothelial and astrocyte co-culture model of the blood-brain barrier utilizing an ultra-thin, nanofabricated silicon nitride membrane. Lab Chip 5(1):74–85. https://doi.org/10.1039/b405713a 26. Bell RD, Winkler EA, Singh I, Sagare AP, Deane R, Wu Z, Holtzman DM, Betsholtz C, Armulik A, Sallstrom J, Berk BC, Zlokovic BV (2012) Apolipoprotein E controls cerebrovascular integrity via cyclophilin A. Nature 485 (7399):512–516. https://doi.org/10.1038/ nature11087 27. Zhao Z, Sagare AP, Ma Q, Halliday MR, Kong P, Kisler K, Winkler EA, Ramanathan A, Kanekiyo T, Bu G, Owens NC, Rege SV, Si G, Ahuja A, Zhu D, Miller CA, Schneider JA, Maeda M, Maeda T, Sugawara T, Ichida JK, Zlokovic BV (2015) Central role for PICALM in amyloid-beta blood-brain barrier transcytosis and clearance. Nat Neurosci 18 (7):978–987. https://doi.org/10.1038/nn. 4025 28. Yuan F, Leunig M, Berk DA, Jain RK (1993) Microvascular permeability of albumin, vascular surface area, and vascular volume measured in human adenocarcinoma LS174T using dorsal chamber in SCID mice. Microvasc Res 45(3):269–289. https://doi.org/10.1006/ mvre.1993.1024 29. Kimura M, Dietrich HH, Huxley VH, Reichner DR, Dacey RG Jr (1993) Measurement of hydraulic conductivity in isolated arterioles of rat brain cortex. Am J Phys 264(6 Pt 2): H1788–H1797. https://doi.org/10.1152/ ajpheart.1993.264.6.H1788 30. Xiang TX, Anderson BD (1994) The relationship between permeant size and permeability in lipid bilayer membranes. J Membr Biol 140 (2):111–122. https://doi.org/10.1007/ bf00232899

Methods in Molecular Biology (2021) 2367: 123–135 DOI 10.1007/7651_2020_315 © Springer Science+Business Media New York 2021 Published online: 11 March 2021

Assessment of Blood-Brain Barrier Permeability Using Miniaturized Fluorescence Microscopy in Freely Moving Rats Jeffrey L. Barr, G. Cristina Brailoiu, Ellen M. Unterwald, and Eugen Brailoiu Abstract We report here the method of visualization of brain microcirculation and assessment of blood-brain barrier (BBB) permeability changes using the miniature integrated fluorescence microscope (i.e., miniscope) technology in awake, freely moving rats. The imaging cannula is implanted in the brain area of interest of anesthetized adult rats. After recovery and habituation, sodium fluorescein, a low-molecular-weight tracer, is injected i.v. Fluorescence intensity in the vicinity of microvessels, as an indicator of BBB permeability, is then recorded in vivo via the miniscope for extended periods of time. The method can be used to assess the changes in BBB permeability produced by pharmacologic agents; in this case, the drug of interest is administered after sodium fluorescein. An increase in the sodium fluorescein extravasation in brain microcirculation demonstrates an increase in BBB permeability. The method described here allows a highresolution visualization of real-time changes in BBB permeability in awake, freely moving rats. Keywords Brain microcirculation, Fluorescence imaging, In vivo BBB permeability, Sodium fluorescein, Vascular extravasation

1

Introduction The blood-brain barrier (BBB) is a complex structure that separates the circulation from the central nervous system; it regulates the transport of molecules in both directions and protects and maintains the homeostasis of neural microenvironment [1]. Endothelial cells of the BBB, connected via tight and adherens junctions, form a tight layer with reduced permeability [2]. BBB dysfunction has been involved in pathogenesis of neurodegenerative disorders such as Alzheimer’s disease, Parkinson’s disease, and multiple sclerosis, or in acute CNS disorders such as stroke, traumatic brain injury, spinal cord injury, and epilepsy [2].

Electronic Supplementary Material: The online version of this chapter (https://doi.org/10.1007/7651_ 2020_315) contains supplementary material, which is available to authorized users.

123

124

Jeffrey L. Barr et al.

Our current understanding of the cellular and molecular mechanisms of BBB function is largely based on in vitro models that assess transendothelial electrical resistance (TEER), permeability to fluorescein-dextran or distribution of tight and adherens junctions in primary or immortalized cerebral microvascular endothelial cells from various species, alone or in co-culture [3]. In vivo investigation of BBB permeability usually consists of assessment of brain extravasation of endogenous tracers (such as albumin, immunoglobulin G, or fibrinogen) or of injected molecules that cannot cross the intact BBB [4–6]. Sodium fluorescein and Evans blue are commonly used as low- and high-molecular-weight tracers; their brain accumulation is assessed ex vivo as an indication of permeability of the barrier [4, 6–8]. Methods used for in vivo BBB imaging in small animals in experimental settings include fluorescence, nuclear, and magnetic resonance imaging [5]. An important tool for the high-resolution dynamic fluorescence imaging of BBB is the two-photon and multiphoton laser scanning microscopy [9, 10]; it allows time-lapse BBB imaging combined with the comparison and correlation with astrocytic and neuronal responses [11, 12]. However, it involves a complex and invasive technique performed in anesthetized animals [11, 12]. Noninvasive fluorescence imaging with Cy5.5 or Cy5.5 conjugated with bovine serum albumin allows the live imaging of deep areas of brain microcirculation and BBB in mice with high sensitivity but limited resolution; the resolution is improved by near-infrared fluorescence imaging (NIRF) [13, 14]. Other imaging techniques used for assessment of BBB function in small animals are nuclear imaging [15] and magnetic resonance imaging [16, 17]. These imaging techniques involve specialized and expensive equipment and are performed in anesthetized animals; a limitation is that anesthetic agents may influence BBB function [18]. There is a need of developing experimental approaches to investigate the BBB function in the living brain. We recently reported the use of miniature integrated fluorescence microscope (i.e., miniscope) technology to assess the changes in BBB permeability [19]. This state-of-the-art technology was developed to assess neuronal activity in vivo using fluorescence probes [20]. Miniscope imaging of brain microcirculation allows the highresolution, real-time visualization of fluorescent tracer extravasation at selected regions of interests (ROIs) in freely moving rats [19].

2

Materials

2.1 Reagents and Supplies for Surgery and Imaging

1. Anesthetics and analgesics: ketamine, xylazine, isoflurane, and meloxicam (see Note 1). 2. Disinfectants: ethanol 70%, chlorhexidine solution.

Use of Miniscope for In Vivo Measurement of BBB Permeability in Awake Rats

125

3. Sterile saline. 4. Lubricant eye ointment. 5. Dental cement. 6. Cyanoacrylate glue gel. 7. Sodium fluorescein: 200 mg/kg i.v. (for imaging). 8. Cotton swabs, gauze sponges. 9. Anchor screws. 10. Syringes (1 and 3 mL) and needles (27G). 2.2 Equipment and Tools for Surgery and Imaging

1. Isoflurane anesthesia system (recommended). 2. Surgical tools: surgical scalpel, scissors, fine sharp forceps. 3. Laboratory balance. 4. Electric hair shaver. 5. Small animal stereotaxic apparatus (such as Kopf, or updated computer-controlled model) with non-rupture ear bars. 6. Dissecting microscope. 7. Handheld drill with small trephine drill bit (for lens cannula) and bits for anchor screws. 8. Temperature-controlled heating pad or heat lamp. 9. Jewelers screwdriver. 10. Doric miniscope setup, cannula (GRIN lens), and focus ring (Doric Lenses Inc, Quebec, Canada; http://doriclenses.com/). 11. Doric Studio software (Doric Lenses Inc).

2.3

Animals

1. Adult rats male or female rats, 300 g. 2. Rats are housed on a 12-h light/dark cycle with free access to standard lab chow and water throughout the study. 3. Animal protocols are approved by the Institutional Animal Care and Use Committee of Temple University.

3

Methods

3.1 Surgery Preparation

1. Prior to procedures, disinfect the surgical area with 70% ethanol. Surgical tools should be sterilized by autoclaving (121  C for 15 min or 131  C for 3 min using steam sterilization) or by cleaning with disinfectant. Between surgeries, instruments are wiped clean with alcohol-soaked sterile gauze and then placed in a hot glass bead sterilizer for 60 s followed by rinsing with cool sterile saline (see Note 2).

126

Jeffrey L. Barr et al.

2. Wear disposable gown, face mask, head cover, closed-toe shoes, shoe covers. 3. Clean hands well using an antibacterial soap (e.g., chlorhexidine or iodophor), and wear surgical gloves. 4. Measure cannula for the desired penetration depth, and use the designed focusing ring. 3.2

Anesthesia

1. Anesthetize rats with ketamine/xylazine or isoflurane. 2. When using a mixture of ketamine and xylazine, weigh the animal, and calculate the appropriate dose for anesthesia. Use a dose of 80 mg ketamine and 10 mg xylazine per kilogram body weight, given intraperitoneally, for adult rats (see Note 3). 3. When using isoflurane, turn on the isoflurane machine at a 1 L/min flow rate of oxygen. Place the animal in the induction chamber, and then increase the percentage of isoflurane until it reaches 5%. Keep the animal in the chamber until a deep level of anesthesia is achieved. 4. Monitor the depth of anesthesia continuously prior to the start of surgery and throughout the surgical procedure (every ~10 min) by inspection of reflexes including the withdrawal reflex when the hind foot pad is pinched, as well as respiratory rate. 5. Once the toe pinch response disappears, breathing is rapid and steady, and there is no response to surgical stimulation, the animal will be considered adequately anesthetized to perform surgery. 6. Meloxicam (1 mg/kg, i.p.) will be administered prior to the start of surgery (after anesthesia, before first incision).

3.3 Stereotaxic Cannulation Surgery

1. Use aseptic techniques. 2. Shave the fur on the top of the head closely in a wide margin to prevent hair contamination. Clean the skin with three alternating chlorhexidine and ethanol scrubs. 3. Place the rat in the stereotaxic apparatus using only non-rupture ear bars. The animal’s head should be flat horizontally with respect to the ear bars (see Note 4). 4. Use small forceps to pull down the lower jaw, slowly move the incisor bar into the mouth until the incisors are in the opening, then gently pull back slightly, and fix the adaptor in place. The last point of fixation, the nose clamp, should be used with very low pressure on the nose. 5. If using isoflurane, carefully move the nose cone attached to the incisor bar forward to cover the nose. Decrease the percentage of isoflurane, and maintain an isoflurane concentration of 2.0–3.0% for the remainder of the procedure.

Use of Miniscope for In Vivo Measurement of BBB Permeability in Awake Rats

127

6. The animal should be placed on circulating water heating pad (37  C). 7. Apply lubricating eye ointment to prevent corneal drying during the surgery. 8. Make an incision in the center of the scalp with a sterile scalpel extending from the lambda to just between the eyes. Use hemostats or clips to pinch off the skin, and keep the incision open. Using several sterile cotton swabs, dry off the exposed skull surface. Cauterize any bleeding areas of the skull. 9. Gently clean the bregma and lambda areas by scraping with the scalpel. 10. Level the head anterior/posterior by measuring the z coordinates of bregma and lambda and adjusting the incisor bar until they become equal. 11. Measure the position of the x and y coordinates of bregma, and calculate the coordinates of the target area, as determined from a stereotaxic brain atlas [21]. Mark the skull at the future craniotomy site. 12. Drill and place 2–3 anchor screws in the skull plates surrounding the craniotomy site to provide extra stability to the mount. 13. Begin craniotomy using a trephine drill bit in a handheld drill (see Note 5). Use a small needle to perforate the edges of the craniotomy, and carefully remove it with fine forceps. Keep the exposed dura moist with sterile phosphate-buffered saline (PBS). 14. Place the cannula onto the microscope fixed to the microscope holder on the stereotaxic arm. Choose the type of cannula from Doric Lenses based on the targeted brain region. 15. Bring the cannula tip to the correct x and y position, and lower it until it touches the exposed dura. Measure the reference point, and cut a small incision in the dura with a 27-G needle held at an angle (see Note 6). 16. Slowly lower (0.6 mm/min) the lens to the desired z coordinate. For example, for cannula implantation into the prefrontal cortex of a 300 g rat, the following stereotaxic coordinates from bregma can be used (AP, 3 mm; ML, 0.5 mm; DV, 2.6 mm) [19, 21]. 17. Once the cannula is implanted, place cyanoacrylate glue gel between the focusing ring and the skull. When the glue is completely dried, apply dental cement to cover the base of the focusing ring and screws and exposed skull (see Note 7). 18. Detach cannula from miniscope, and move the holder arm away from the animal.

128

Jeffrey L. Barr et al.

3.4 Post-surgery Care and Monitoring

1. Inject warm sterile saline (~37  C, 10 mL/kg, i.p.) to avoid dehydration of the animal after the surgery and meloxicam (1 mg/kg, s.c.). 2. Keep the rat warm (either on a temperature-controlled heat pad or under a heat lamp) until it fully recovers. Post-surgical monitoring should be performed every 10 min until fully awake and ambulatory (see Note 8). 3. Provide rats with contact stimulation, and vary their position every 15 min to improve respiration and heat generation. 4. Once the animal fully recovers, return the animal to a clean cage (single-housed) with a DietGel pack in the cage for easy access to food. 5. Administer additional analgesic (e.g., meloxicam at a dose of 1 mg/kg, s.c.) on the day after the surgery. 6. Closely monitor the recovery of the animal for at least 1 week, assessing any signs of distress, such as piloerection, lack of grooming, reduced locomotion, wound scratching, or inflammation of the surgical site.

3.5 Habituation to the Miniscope

1. After 1 week of post-surgical recovery, habituate rats to the miniscope by attaching the dummy miniscope to the cannula and allowing the animal to freely explore for 10 min. The dummy miniscope snaps into place onto the imaging cannula during minimal manual restraint. 2. Increase the time by 5 min each day until the full time needed for imaging is obtained. 3. Imaging sessions can be carried out 2 weeks after cannula implantation (Fig. 1 and Supplementary Video 1). 4. Monitor animals continuously during the habituation sessions.

3.6 Imaging Session Preparation

1. Weigh each rat before injections. Record the body weight and the agent volume to be administered for each animal. 2. Prior to tail vein injection of tracer, warm the rat for 5–10 min to dilate the veins (e.g., by using a warm water circulating pad or overhead heat lamp or by soaking the tail in warm water (30–35  C) to cause vasodilation). 3. Rats can be lightly anesthetized or restrained using a commercially available restraint device (see Note 9). 4. Position the animal on its side. Insert the needle (27 gauge) parallel to the tail, bevel facing up, into the vein toward the direction of the head. 5. Slowly inject sodium fluorescein (Na-F) (2% solution in saline, 0.5 mL/kg, filter sterilized, filter size, 0.2 μm). If there is any

Use of Miniscope for In Vivo Measurement of BBB Permeability in Awake Rats

129

Fig. 1 Rat with implanted cannula (image and video). Illustration of rat bearing cannula implanted into the prefrontal cortex 2 weeks earlier

resistance or a blister appears on the tail, the needle should be removed and reinserted above the first site. 6. Remove the needle, and apply gentle compression until any bleeding has stopped. 7. Connect the miniscope to the previously implanted imaging cannula. 8. If the animal was anesthetized, allow full recovery before imaging. 3.7 Fluorescence Recording Using Miniscope

1. Allow 15–30 min for equilibration after injection of sodium fluorescein. 2. Visualize sodium fluorescein (excitation/emission 488/520 nm) in microvessels using the minimum exposure time (100 ms) and minimum intensity (50–75% intensity light or less) to avoid photobleaching. 3. Record one frame every 5 min; a rapid conversion to pseudocolor will easily indicate if the preparation is stable to begin the experiment. 4. Imaging experiments can be performed up to 6 weeks after cannulation. After this, the imaging cannula becomes cloudy due to the deposition of debris on the lens (see Note 10; Fig. 2).

130

Jeffrey L. Barr et al.

Fig. 2 Imaging cannula. Comparison of a new cannula versus one that was used for 6 weeks. Circled in the center is the imaging cannula from the regular (a) or eFocus miniscope (b); the new cannula is seen as a shiny surface, while after 6 weeks, it becomes cloudy 3.8 Image Analysis of Fluorescence Recordings

1. Draw regions of interest (ROIs) in close proximity of microvessels (to assess sodium fluorescein extravasation) maintaining the same distance from the center of the blood vessel (see Note 11; Fig. 3). 2. Visualize, record, and analyze post-acquisition the fluorescence in the same ROIs over time using Doric Studio software (Doric Lenses Inc) or another software such as Image J (see Note 12, Figs. 4 and 5). 3. Data can be expressed as mean fluorescence intensity (SEM) for several ROIs (e.g., 10 ROIs), normalized to control baseline fluorescence measured immediately prior to injection of saline or test agent in the same subject [19].

Use of Miniscope for In Vivo Measurement of BBB Permeability in Awake Rats

131

Fig. 3 Diagram indicating position of the region of interest (ROI). Example of a ROI drawn in the proximity of microvessel; its distance from the center of the blood vessel is indicated. To assess sodium fluorescein extravasation as an indicator of BBB permeability, the fluorescence intensity of multiple ROIs of the same size and at the same distance from the center of the blood vessel can be determined and averaged

4

Notes 1. Ketamine should be handled according to relevant rules of the host institution. 2. A new set of sterilized instruments should be used as recommended by the IACUC. 3. Rats anesthetized with a mixture of ketamine (80 mg/kg) and xylazine (10 mg/kg) should reach surgical anesthesia (lack of response to nociceptive stimuli) within 5–10 min. 4. When the rat is placed in the stereotaxic apparatus, the head should be free to move vertically but not laterally. 5. Performing the craniotomy with a handheld drill makes the implantation of a lens easier with less risk of breaking the lens. Do not drill completely through the bone in order to avoid injury to the surface of the brain. 6. When measuring the reference point, a clean brain surface, without bleeding, is needed. 7. Allow the dental cement to completely dry before removing the animal from the apparatus. 8. Post-surgical monitoring includes observation of respiratory rate/quality, mucous membrane color, activity, anesthesia depth, and incisional integrity including any leakage or bleeding. 9. The duration of the restraint should be kept to a minimum. 10. The imaging cannula can be used up to 6 weeks (Fig. 2). 11. Maintain the same area of the ROI and the same distance from the center of the blood vessel (Fig. 3). 12. Sodium fluorescein extravasation, as an indicator of BBB permeability, is measured immediately prior to (i.e., baseline) and

132

Jeffrey L. Barr et al.

Fig. 4 Illustration of brain microcirculation imaged with miniscope after injection of sodium fluorescein (Na-fluorescein). Example of Na-fluorescein fluorescence visualized with miniscope in rat prefrontal cortex (a) and pseudocolor images processed with Image J (b) in rat injected with control saline (min 0) and 30 min after saline. An example of region of interest (ROI) is indicated in each panel. (c) Comparison of the Na-fluorescein extravasation measured in the indicated ROI at the two time points after saline injection (0 and 30 min)

again 3–30 min following saline (Fig. 4) or the experimental agent using a within-subjects design [19]. The effect of cocaine (20 mg/kg, i.p.) which increases BBB permeability [19, 22] is illustrated in Fig. 5.

Use of Miniscope for In Vivo Measurement of BBB Permeability in Awake Rats

133

Fig. 5 Cocaine induces an increase in sodium fluorescein (Na-fluorescein) extravasation in the rat brain. Examples of Na-fluorescein fluorescence visualized with miniscope in rat prefrontal cortex (a) and pseudocolor images (b) before (control cocaine min 0) and 30 min after injection of cocaine (20 mg/kg, i.p.). An example of region of interest (ROI) is indicated in each panel. (c) Comparison of the Na-fluorescein extravasation measured in the indicated ROI at the two time points after cocaine injection (0 and 30 min)

134

Jeffrey L. Barr et al.

Acknowledgments This work was supported by the National Institutes of Health grant P30DA013429. References 1. Abbott NJ, Patabendige AA, Dolman DE, Yusof SR, Begley DJ (2010) Structure and function of the blood-brain barrier. Neurobiol Dis 37(1):13–25. https://doi.org/10.1016/j. nbd.2009.07.030 2. Sweeney MD, Zhao Z, Montagne A, Nelson AR, Zlokovic BV (2019) Blood-brain barrier: from physiology to disease and back. Physiol Rev 99(1):21–78. https://doi.org/10.1152/ physrev.00050.2017 3. Deli MA, Abraham CS, Kataoka Y, Niwa M (2005) Permeability studies on in vitro bloodbrain barrier models: physiology, pathology, and pharmacology. Cell Mol Neurobiol 25 (1):59–127 4. Nag S (2003) Blood-brain barrier permeability using tracers and immunohistochemistry. Methods Mol Med 89:133–144. https://doi. org/10.1385/1-59259-419-0:133 5. Wunder A, Schoknecht K, Stanimirovic DB, Prager O, Chassidim Y (2012) Imaging blood-brain barrier dysfunction in animal disease models. Epilepsia 53(Suppl 6):14–21. https://doi.org/10.1111/j.1528-1167.2012. 03698.x 6. Yen LF, Wei VC, Kuo EY, Lai TW (2013) Distinct patterns of cerebral extravasation by Evans blue and sodium fluorescein in rats. PLoS One 8(7):e68595. https://doi.org/10. 1371/journal.pone.0068595 7. Brailoiu E, Barlow CL, Ramirez SH, Abood ME, Brailoiu GC (2018) Effects of plateletactivating factor on brain microvascular endothelial cells. Neuroscience 377:105–113. https://doi.org/10.1016/j.neuroscience. 2018.02.039 8. Leo LM, Familusi B, Hoang M, Smith R, Lindenau K, Sporici KT, Brailoiu E, Abood ME, Brailoiu GC (2019) GPR55-mediated effects on brain microvascular endothelial cells and the blood-brain barrier. Neuroscience 414:88–98. https://doi.org/10.1016/j.neuro science.2019.06.039 9. Nishimura N, Schaffer CB, Friedman B, Tsai PS, Lyden PD, Kleinfeld D (2006) Targeted insult to subsurface cortical blood vessels using ultrashort laser pulses: three models of stroke. Nat Methods 3(2):99–108. https://doi.org/ 10.1038/nmeth844

10. Prager O, Chassidim Y, Klein C, Levi H, Shelef I, Friedman A (2010) Dynamic in vivo imaging of cerebral blood flow and blood-brain barrier permeability. NeuroImage 49 (1):337–344. https://doi.org/10.1016/j. neuroimage.2009.08.009 11. Zhang S, Murphy TH (2007) Imaging the impact of cortical microcirculation on synaptic structure and sensory-evoked hemodynamic responses in vivo. PLoS Biol 5(5):e119. https://doi.org/10.1371/journal.pbio. 0050119 12. Takano T, Han X, Deane R, Zlokovic B, Nedergaard M (2007) Two-photon imaging of astrocytic Ca2+ signaling and the microvasculature in experimental mice models of Alzheimer’s disease. Ann N Y Acad Sci 1097:40–50. https://doi.org/10.1196/ annals.1379.004 13. Abulrob A, Brunette E, Slinn J, Baumann E, Stanimirovic D (2008) Dynamic analysis of the blood-brain barrier disruption in experimental stroke using time domain in vivo fluorescence imaging. Mol Imaging 7(6):248–262 14. Piper S, Bahmani P, Klohs J, Bourayou R, Brunecker P, Muller J, Harhausen D, Lindauer U, Dirnagl U, Steinbrink J, Wunder A (2010) Non-invasive surface-stripping for epifluorescence small animal imaging. Biomed Opt Express 1(1):97–105. https://doi.org/ 10.1364/BOE.1.000097 15. Rowland DJ, Cherry SR (2008) Small-animal preclinical nuclear medicine instrumentation and methodology. Semin Nucl Med 38 (3):209–222. https://doi.org/10.1053/j. semnuclmed.2008.01.004 16. Nagaraja TN, Ewing JR, Karki K, Jacobs PE, Divine GW, Fenstermacher JD, Patlak CS, Knight RA (2011) MRI and quantitative autoradiographic studies following bolus injections of unlabeled and (14)C-labeled gadoliniumdiethylenetriaminepentaacetic acid in a rat model of stroke yield similar distribution volumes and blood-to-brain influx rate constants. NMR Biomed 24(5):547–558. https://doi.org/10.1002/nbm.1625 17. Jelescu IO, Leppert IR, Narayanan S, Araujo D, Arnold DL, Pike GB (2011) Dualtemporal resolution dynamic contrast-

Use of Miniscope for In Vivo Measurement of BBB Permeability in Awake Rats enhanced MRI protocol for blood-brain barrier permeability measurement in enhancing multiple sclerosis lesions. Journal of magnetic resonance imaging 33(6):1291–1300. https:// doi.org/10.1002/jmri.22565 18. Sharma HS, Muresanu DF, Nozari A, Castellani RJ, Dey PK, Wiklund L, Sharma A (2019) Anesthetics influence concussive head injury induced blood-brain barrier breakdown, brain edema formation, cerebral blood flow, serotonin levels, brain pathology and functional outcome. Int Rev Neurobiol 146:45–81. https:// doi.org/10.1016/bs.irn.2019.06.006 19. Barr JL, Brailoiu GC, Abood ME, Rawls SM, Unterwald EM, Brailoiu E (2020) Acute cocaine administration alters permeability of blood-brain barrier in freely-moving rats-evidence using miniaturized fluorescence

135

microscopy. Drug Alcohol Depend 206:107637. https://doi.org/10.1016/j.dru galcdep.2019.107637 20. Ghosh KK, Burns LD, Cocker ED, Nimmerjahn A, Ziv Y, Gamal AE, Schnitzer MJ (2011) Miniaturized integration of a fluorescence microscope. Nat Methods 8 (10):871–878. https://doi.org/10.1038/ nmeth.1694 21. Paxinos G, Watson C (2013) The rat brain in stereotaxic coordinates, 7th edn. Academic Press 22. Sharma HS, Muresanu D, Sharma A, Patnaik R (2009) Cocaine-induced breakdown of the blood-brain barrier and neurotoxicity. Int Rev Neurobiol 88:297–334. https://doi.org/10. 1016/S0074-7742(09)88011-2

Methods in Molecular Biology (2021) 2367: 137–148 DOI 10.1007/7651_2020_345 © Springer Science+Business Media, LLC 2021 Published online: 19 January 2021

Measurement of Lung Vessel and Epithelial Permeability In Vivo with Evans Blue Prestina Smith, Lauren A. Jeffers, and Michael Koval Abstract Lung fluid balance is maintained in part by the barriers formed by the pulmonary microvasculature and alveolar epithelium. Failure of either of these barriers leads to pulmonary edema, which limits lung function and exacerbates the severity of acute lung injury. Here we describe a method using Evans Blue dye to simultaneously measure the function of vascular and epithelial barriers of murine lungs in vivo. Key words Acute respiratory distress syndrome, Alveolar epithelium, Lung barrier function, Pulmonary edema, Tight junctions, Vascular endothelium

1

Introduction In order to efficiently mediate gas exchange, the lung needs to efficiently control fluid balance. Injury and inflammation impair the control of lung fluid, resulting in pulmonary edema which, when extensive, exacerbates lung injury and causes acute respiratory distress syndrome (ARDS) [1]. Control of the lung air/liquid barrier is predominantly mediated by two distinct systems, the pulmonary microcirculation [2, 3] and the alveolar epithelium [4], failure of which can lead to interstitial fluid accumulation (tissue edema) and airspace flooding, respectively (Fig. 1). Tissue edema and airspace flooding both have the capacity to impair gas exchange and exacerbate injury. Understanding the relative contributions of the vascular endothelial and alveolar epithelial barriers in regulating lung fluid is critical to identifying effective therapeutic approaches to the treatment of ARDS. It has long been appreciated that Evans Blue dye (T-1824) preferentially binds to serum albumin [6], making it an effective marker for the ability of albumin to extravasate across barriers and accumulate in tissues. Albumin is a 68 kDa, native serum protein that has been found to penetrate barriers and accumulate in airspaces in response to lung injury (e.g., [7]), which underscores its utility as a marker for lung barrier failure. Albumin permeability also is a particularly useful marker in that it can cross endothelial

137

138

Prestina Smith et al.

Fig. 1 Diagram depicting a cross-section though a normal lung alveolus comprised of an epithelial sac surrounded by blood vessels. Note that the transit of fluid-borne molecules from the vasculature must traverse across the vascular endothelial barrier though the interstitial tissue and across the alveolar epithelial barrier. (Reproduced from [5] with permission)

and epithelial barriers by both the paracellular route through tight junctions [8] and the transcellular route through transcytosis [9– 11]. In contrast to direct measurement of albumin in different lung compartments by ELISA or immunoblot, Evans Blue is easily measured by absorbance spectroscopy and is highly sensitive [12]. This is particularly critical for measurement of dye accumulation in the interstitium. Here we describe a protocol that enables the accumulation of Evans Blue in the airspace and interstitial compartments to be simultaneously determined, in order to measure the relative impact of injury on lung endothelial and epithelial barriers.

2

Materials

2.1 Evans Blue Injection

1. Dulbecco’s Phosphate Buffered Saline (DPBS) without Ca2+/ Mg2+ (Corning/Mediatech #21-031-CV, Manassas, VA). 2. Evans Blue: Make a 0.5% solution in DPBS by adding 0.05 g of Evans Blue (MilliporeSigma E2129, St. Louis, MO) to 10 ml of DPBS (w/o Ca2+/Mg2+), then filter sterilize. Store at room temperature protected from light.

Measurement of Lung Vessel and Epithelial Permeability In Vivo with Evans Blue

139

3. Sedative/Anesthetic cocktail (20% Ketamine +5% xylazine solution): Add 1 ml 100 mg/ml ketamine (Mylan Institutional NDC 67457-108-10, Galway, Ireland) and 0.25 ml 20 mg/ml xylazine (Zoetis Inc. NDC 59399-110-20, Kalamazoo, MI) to 3.75 ml DPBS without Ca2+/Mg2+ to produce 5 ml total solution. 4. Isoflurane (Piramal Enterprises Limited NDC 66794-017-25, Telangana, India). 5. 25G
5/8 needles (BD PrecisionGlide #305122, Franklin Lakes, NJ) and 30G
1 needles (BD PrecisionGlide #305128) (see Note 1). 6. 1 ml syringes (BD #309659). 7. Cotton Gauze Sponges, 5
5 cm (FisherBrand #22-362-178, Pittsburgh, PA). 8. Heat lamp. 9. Small animal heating pad. 2.2

Tissue Harvest

1. Animal surgical instruments: dissecting scissors, fine tip tweezers, smooth and rat tooth forceps, dissection board. 2. Portable balance to weigh animals (Mettler Toledo PL6000-S or equivalent). 3. Mouse trachea tube (DWK Kimble Kontes Brand Microflex Syringe Needles-Blunt, Fisher Scientific #K868280-2001). 4. 5-0 Silk Suture (Ethicon #K870H, Somerville, NJ). 5. 21G
1 ½ needles (BD PrecisionGlide #305167). 6. 10 ml syringe (BD #302995). 7. 1.5 ml conical microfuge tubes (Eppendorf, # 022364111, Enfield, CT). Each mouse requires 3 collection tubes each for blood/serum, bronchoalveolar lavage (BAL) fluid, and lung tissue. 8. Heparin (Aurobindo Pharma Limited NDC 63739–920-25, Memphis, TN) stock concentration 1,000 USP: Used to coat needle and syringe for blood collection. 9. Formamide (Electron Microscopy Sciences #15745, Hatfield, PA). Add 250 μl per tube for extraction of lung tissue. 10. DPBS with Ca2+/Mg2+ (Corning/Mediatech #21-030-CV). 11. Lavage solution: DPBS with Ca2+/Mg2+ containing 1:200200 mM phenylmethylsulfonyl fluoride (PMSF, MilliporeSigma, # P7626) and 1:500 1 M NaF (MilliporeSigma #S7920): Prepare 500 μl per mouse and keep lavage fluid on ice in an ice bucket (see Note 2).

140

2.3

Prestina Smith et al.

Tissue Analysis

1. Heating block (Fisher Scientific Dry Bath Incubator #11–718 or equivalent). 2. Refrigerated microfuge (Eppendorf 5415 R or equivalent). 3. Microplate reader (Biotek Synergy H1 Multimode Plate Reader or equivalent).

3

Methods

3.1 Tail Vein Injection of Evans Blue

1. Prepare 0.5% Evans Blue solution. 2. Place mice in a cage under the heat lamp. Warm mice for 5–10 min in order to dilate blood vessels (see Note 3). 3. Weigh each mouse and intraperitoneally (IP) inject 10 μl/g body weight + 10 μl ketamine/xylazine anesthetic cocktail (see Note 4). 4. Place the mice back in the cage and allow anesthetic to take effect (see Note 5). 5. Place mouse on a platform so that the mouse is at a comfortable height and the tail hangs over the edge. 6. Rotate the tail to locate the lateral tail vein and potential injection site. Make sure the vein is facing upward (Fig. 2). 7. Clean injection area with warm alcohol pad. 8. Fill syringe with 400 μl Evans Blue solution making sure to avoid air bubbles (see Note 6). 9. Hold the tail with the nondominant hand while stretching the tail so that the injection site is visible and the vein is in a straight line. 10. Insert 30G needle at a 10–15 angle, bevel up, into the injection site. Advance needle towards the head keeping the needle and syringe parallel to the tail. The needle should be visibly in the tail vein (Fig. 2). 11. Slowly inject 200 μl of dye into the tail vein of the mouse. If you have correct placement of the needle the plunger will advance with ease (see Note 6). 12. Place mouse on heating pad and wait 1 h (see Note 7). Evans Blue will clearly dye the nose and footpads of treated mice (Fig. 3).

3.2

Serum Collection

1. Rinse needle (21G) and 1 ml syringe with heparin in order to coat both needle and syringe (see Note 8). 2. Sacrifice the mouse using an approved injectable anesthetic overdose procedure. A 1 ml IP injection 1 ml of 20% Ketamine +5% xylazine solution is sufficient (see Note 9).

Measurement of Lung Vessel and Epithelial Permeability In Vivo with Evans Blue

141

Fig. 2 Tail vein injection. (a, b) Diagram showing mouse tail vasculature. Branch vessels extend from the artery to the veins. Red arrows, artery; blue arrows, vein; green arrows, branch vessels. (c–f) Procedure for tail vein injection. (c) The outer tube is grasped by the first and second fingers. The third finger is placed under the inner cylinder. Place the needle on the surface of the tail in parallel (d) and insert it carefully (e). Once the needle tip is under the skin, pull back the syringe slightly during insertion to confirm that blood flows back to ensure that a vein is penetrated (f, arrow). (Reproduced from [13] with permission)

3. Once unresponsive, pin mouse legs down on dissection board. Immobilize the head by placing a suture loop around the front incisors and pinning the loop down to the dissection board (see Note 10).

142

Prestina Smith et al.

Fig. 3 Appearance of Evans Blue in mice and murine lungs. (a, b) Mice that were either untreated (a) or intravenously injected with Evans Blue (b). Sixty minutes after injection, the treated mice show clear blue labeling of the nose and paws. (c–e). Representative images of lung tissue from mice that were either untreated (c), or after Evans Blue tail vein injection showing control (d) or alcohol fed and endotoxin treatment to induce acute lung injury (e). Uninjured mice show little tissue accumulation (arrow) of Evans Blue as compared with injured mice

4. Wet the abdomen with water or ethanol to prevent fur from entering the abdominal cavity and contaminating tissue. Cut open the skin and peritoneum along the midline. 5. Cut though the ribcage and open chest cavity so that the bottom of the heart is exposed. 6. Puncture the left ventricle with the 21G needle attached to the heparinized 1 ml syringe. Slowly draw plunger back into syringe to start collecting blood. Collect a minimum of 250 μl blood/mouse. 7. Evacuate blood into collection tube and place on ice (see Note 11). 8. Centrifuge blood at 3000
g at 4  C for 15 min. Collect supernatants (serum) and transfer to a new tube. Freeze at 20  C (see Note 12).

Measurement of Lung Vessel and Epithelial Permeability In Vivo with Evans Blue

3.3 BAL Fluid Collection

143

1. Cut away skin on the neck so that the area near the trachea is exposed. Carefully snip away thin tissue covering the trachea (see Note 13). 2. Turn the mouse so that the head is closer to you. 3. With the help of forceps, lay a string of surgical sutures behind the trachea. 4. Make a small nick at a 45 angle on the top surface of the trachea nearest the larynx (see Note 14). 5. Slide the 20G trachea tube into the nick and stop right above the ribcage. Ensure the surgical string is at least 2–3 mm above the bottom tip of the trachea tube. 6. Secure the needle in place by tying a knot with the sutures around it. 7. Remove any air in the lungs by evacuating at least 1 ml of air from the lung using an empty 1 ml syringe. 8. Quickly replace the empty syringe with one filled with 500 μl of lavage fluid and slowly fill lungs with the fluid. Rinse the lungs twice with the same lavage fluid, then pull the plunger out to retrieve the final volume (see Note 15). 9. Collect the fluid in centrifuge tubes and place on ice. 10. Centrifuge BAL at 3000
g at 4  C for 15 min. Collect supernatants and transfer to a new tube. Freeze at 20  C (see Note 12).

3.4 Lung Tissue Collection

1. Rotate the mouse back to the original position with the feet closest to you. 2. Snip the aorta below the diaphragm so that blood can easily move from the perfused lungs (see Note 16). 3. Take a DPBS with Ca2+/Mg2+-filled 10 ml syringe attached to a 25G needle and insert the needle into the right ventricle, aiming in the direction of the pulmonary artery. Gently push the plunger of the syringe to perfuse the lungs with 5 ml of DPBS (see Note 17). 4. Carefully dissect out the lungs from the thoracic cavity by grabbing the trachea with forceps then snipping down behind the lungs while gently pulling the lungs away. 5. Once out, rinse the exterior of the lungs with DPBS with Ca2+/Mg2 to remove any clotted exterior blood and cut away extrapulmonary tissue (e.g., heart and vasculature) and remove cartilaginous tissue (trachea and large bronchi). Lungs with tissue accumulation of Evans Blue are visibly stained (Fig. 3c–e).

144

Prestina Smith et al.

6. Place half the isolated lung tissue in a centrifuge tube containing 250 μl of formamide on ice. Weigh lung samples and record for later analysis (see Note 18). 7. Incubate at 55  C in a heating block for 48 h. 8. Centrifuge lung tissue at 3000
g at 4  C for 15 min. Collect supernatants and transfer to a new tube. 3.5 Evans Blue Measurement and Analysis

1. Aliquot 100 μl/well of samples from serum (diluted 1:10 in PBS), undiluted BAL Fluid, and lung tissue extract in a 96-well plate (see Note 19). 2. Measure absorbance of samples and standards at 620 nm and 740 nm using a microplate reader (see Note 20). 3. Correct the A620 readings for turbidity in BAL fluid and lung tissue using the correction factor y ¼ 1.193x + 0.007 where x is A740 and A620 corrected ¼ yA620 [12]. Use standards to get absolute Evans Blue concentrations in μg/ml. 4. Divide BAL fluid Evans Blue concentration by serum Evans Blue concentration to normalize and account for variability in the tail vein injections (see Note 21). 5. Divide lung tissue Evans Blue concentration by lung weights and then by serum Evans Blue concentration (see Note 22).

4

Notes 1. 30G needles are used for tail vein injection but 25G needles are used for sedation, since 30 G needles are too small to quickly sedate mice at the volumes required. 2. Pre-cooling lavage fluid on ice before BAL extraction significantly increases the yield and helps inhibit protease activity when harvesting BAL. 3. Make sure that the heat lamp is not too close to the cage by placing your hand near the bottom of the cage and holding for 30 s. The lamp should be no closer than 20–30 cm from the mice. If the heat is uncomfortable to you the lamp is too close and may cause the mice to overheat. 4. This anesthetic guideline is for mice weighing 25–35 g. For mice smaller than 25 g, use 10 μl/g body weight ketamine/ xylazine. For mice larger than 35 g, use 10 μl/g body weight + 15 μl ketamine/xylazine. Alternatively, Evans Blue injection can be done on mice without anesthetic using a mouse restraint device (Red Tailveiner Restrainer Braintree Scientific Inc. #TV-RED 150-STD, Braintree, MA). Note that sedated and nonsedated mice will give different Evans

Measurement of Lung Vessel and Epithelial Permeability In Vivo with Evans Blue

145

Blue permeability values due to differences in heart rate, breathing rate, and stimulation of the sympathetic system. 5. Mice usually are nonresponsive to a foot pinch after 5 min as an indication that they are anesthetized. 6. Although the syringe is filled with 400 μl Evans Blue solution, only 200 μl is administered. Having an excess of dye solution in the syringe enables the amount injected to be more accurately monitored, especially if the tail vein is initially missed. If it is difficult to advance the plunger the needle is not in the tail vein. The entire mouse should turn blue within 1 min, most notably in the pads of the feet, the nose, and the ears. 7. The heating pad step is only required if anesthetic was used. The heat keeps mice warm as they recover to avoid a drop in body temperature due to the anesthetic. If mice were not anesthetized, place the mouse back into the cage for 1 h. 8. After 1 h mice are ready for tissue harvest. Harvest in the following order: (1) blood, (2) BAL fluid, (3) lung tissue. 9. Overdose with ketamine/xylazine is the preferred method of euthanization since CO2 asphyxiation stops the heart from beating and reduces recovery of blood during collection. Harvest blood immediately after sacrifice for maximum collection. Do not use cervical dislocation, since this can tear the trachea rendering mice unable for BAL collection. 10. The head should be tilted back fully with the neck fully exposed. 11. It can be difficult to distinguish serum from red blood cells because they will be heavily dyed with Evans Blue. By collecting at least 200 μl of whole blood, the top 60 μl supernatant can be isolated without disrupting the cellular layer on the bottom. 12. Serum and centrifuged BAL fluid are stored frozen prior to analysis since the lung tissue requires 48 h processing time. 13. The trachea is beneath the salivary glands. Glands can be carefully cut away and gently pulled apart to uncover the trachea. 14. The nick should only cut the top surface of the trachea and should be just large enough to accommodate the 20G trachea tube. The cut should open less than half of the diameter of the trachea. 15. The lungs should clearly inflate with the addition of fluid. Typical fluid recovery is 300–350 μl of lavage fluid. This provides enough material for duplicate absorbance measurements and protein determination by the Bicinchoninic Acid (BCA) assay (Sigma-Aldrich #BCA1-1KT).

146

Prestina Smith et al.

16. Snipping the aorta reduces the pressure needed when perfusing with PBS into the right ventricle, avoiding potential damage to the lung microvasculature. 17. Lungs should turn white (with patches of blue dye) in color with good perfusion. It is imperative that the lung is properly perfused so that dye from the blood will not interfere with the tissue analysis (Fig. 3c–e). 18. Before adding lung tissue, weigh the tubes containing formamide solution alone. This enables net lung weight to be calculated and used for data analysis. Be consistent in analyzing the right or the left lung tissue/lobes. The remaining lung tissue can be snap frozen in liquid nitrogen and used for other analysis (e.g., immunoblot, Q-PCR). 19. Serum must be diluted 1:10 before analyzing to avoid crosstalk with other serum components. BAL and lung tissue extract should not be diluted. When possible, measure absorbance from duplicate or triplicate technical replicates. 20. Standards are made from serial dilutions of Evans Blue stock into DPBS with Ca2+/Mg2+ (for serum and BAL fluid) or formamide (for lung tissue samples). 21. In this example (Fig. 4), mice were given either a control or alcohol diet for 8 weeks and then challenged by either IT or IP lipopolysaccharide (LPS; endotoxin) as a sterile inflammatory challenge [5]. Alcohol rendered the airspaces prone to flooding even in the absence of an additional insult. Interestingly, LPS decreased Evans Blue permeability into the airspaces, most likely due to an increase in Transforming Growth Factor (TGF)-alpha dependent stimulation of Epidermal Growth Factor (EGF) receptors [14]. IT injury, but not IP injury, of alcohol-fed mice also resulted in failure of the endothelial barrier as indicated by the accumulation of Evans Blue in the interstitial space, consistent with increased severity of direct vs. indirect lung injury on lung fluid balance. 22. Assessing barrier function using Evans Blue dye tracks albumin, which is a 68 kDa protein. While this approach offers many advantages, it represents one parameter when considering the effects of injury on lung fluid balance. For instance, macromolecule permeability may not be impacted under conditions where fluid balance is altered by the effects of injury on ion homeostasis [15].

Measurement of Lung Vessel and Epithelial Permeability In Vivo with Evans Blue

147

Fig. 4 Effect of alcohol consumption and lipopolysaccharide on Evans Blue accumulation in lung airspaces and tissue. Mice received an IP (a, c) or IT (b, d) treatment with either vehicle control or 5 mg/kg LPS in PBS. After 24 h the mice were administered Evans Blue by tail vein injection and allowed to recover for 2 h. Evans Blue dye was first collected by bronchoalveolar lavage (BAL) (a, b). Lung tissue was then harvested after BAL fluid collection and the right ventricle of the heart was perfused with PBS. Evans Blue dye was extracted from lung tissue by incubating in formamide at 55  C for 48 h. Evans Blue concentration was analyzed via spectrophotometry (620 nm). Results were corrected for the presence of heme and normalized to Evans Blue serum levels and lung tissue weight. (a, b) Alcohol-feeding alone increased Evans blue levels in BAL fluid, suggesting that alcohol promotes alveolar epithelial barrier dysfunction (n ¼ 6–9, ****p < 0.0001; **p ¼ 0.0021). (c, d) Tissue Evans Blue dye content was only significantly increased in alcohol-fed mice that were given an IT administration of LPS. There was no significant change in any other groups. n ¼ 4–8, *p ¼ 0.019. Values reported as mean  standard deviation. (Reproduced from [5] with permission)

Acknowledgments Supported by a Ford Foundation Fellowship (PS), NIH grants R01-AA025854 and R01-HL137112 (MK), and F31-HL149323 (LAJ). Experiments were performed in accordance with the National Institutes of Health Guidelines for the Use of Laboratory Animals guidelines and were approved by the Institutional Animal Care and Use Committee at Emory University School of Medicine.

148

Prestina Smith et al.

References 1. Matthay MA, Zemans RL (2011) The acute respiratory distress syndrome: pathogenesis and treatment. Annu Rev Pathol 6:147–163. https://doi.org/10.1146/annurev-pathol011110-130158 2. Mehta D, Bhattacharya J, Matthay MA, Malik AB (2004) Integrated control of lung fluid balance. Am J Physiol Lung Cell Mol Physiol 287(6):L1081–L1090 3. Komarova YA, Kruse K, Mehta D, Malik AB (2017) Protein interactions at endothelial junctions and signaling mechanisms regulating endothelial permeability. Circ Res 120 (1):179–206. https://doi.org/10.1161/ CIRCRESAHA.116.306534 4. Koval M (2017) Junctional interplay in lung epithelial barrier function. In: Sidhaye VK, Koval M (eds) Lung epithelial biology in the pathogenesis of pulmonary disease. Academic Press, Oxford, pp 1–20 5. Smith P, Jeffers LA, Koval M (2019) Effects of different routes of endotoxin injury on barrier function in alcoholic lung syndrome. Alcohol 80:81–89. https://doi.org/10.1016/j.alco hol.2018.08.007 6. Rawson RA (1943) The binding of T-1824 and structurally related diazo dyes by the plasma proteins. Am J Physiol 138(5):708–717. https://doi.org/10.1152/ajplegacy.1943. 138.5.708 7. Bosmann M, Grailer JJ, Zhu K, Matthay MA, Sarma JV, Zetoune FS, Ward PA (2012) Antiinflammatory effects of beta2 adrenergic receptor agonists in experimental acute lung injury. FASEB J 26(5):2137–2144. https://doi.org/ 10.1096/fj.11-201640 8. Schlingmann B, Molina SA, Koval M (2015) Claudins: gatekeepers of lung epithelial

function. Semin Cell Dev Biol 42:47–57. https://doi.org/10.1016/j.semcdb.2015.04. 009 9. Sleep D, Cameron J, Evans LR (2013) Albumin as a versatile platform for drug half-life extension. Biochim Biophys Acta 1830 (12):5526–5534. https://doi.org/10.1016/j. bbagen.2013.04.023 10. Martins JP, Kennedy PJ, Santos HA, Barrias C, Sarmento B (2016) A comprehensive review of the neonatal Fc receptor and its application in drug delivery. Pharmacol Ther 161:22–39. https://doi.org/10.1016/j.pharmthera.2016. 03.007 11. Stewart T, Koval WT, Molina SA, Bock SM, Lillard JW Jr, Ross RF, Desai TA, Koval M (2017) Calibrated flux measurements reveal a nanostructure-stimulated transcytotic pathway. Exp Cell Res 355(2):153–161. https://doi. org/10.1016/j.yexcr.2017.03.065 12. Moitra J, Sammani S, Garcia JG (2007) Re-evaluation of Evans blue dye as a marker of albumin clearance in murine models of acute lung injury. Transl Res 150 (4):253–265. https://doi.org/10.1016/j.trsl. 2007.03.013 13. Hatakeyama S, Yamamoto H, Ohyama C (2010) Tumor formation assays. Methods Enzymol 479:397–411. https://doi.org/10. 1016/S0076-6879(10)79023-6 14. Koff JL, Shao MX, Kim S, Ueki IF, Nadel JA (2006) Pseudomonas lipopolysaccharide accelerates wound repair via activation of a novel epithelial cell signaling cascade. J Immunol 177(12):8693–8700 15. Eaton DC, Helms MN, Koval M, Bao HF, Jain L (2009) The contribution of epithelial sodium channels to alveolar function in health and disease. Annu Rev Physiol 71:403–423

Methods in Molecular Biology (2021) 2367: 149–163 DOI 10.1007/7651_2021_365 © Springer Science+Business Media, LLC 2021 Published online: 09 May 2021

Measurement of Airway Epithelial Permeability: Methods and Protocols Hasan Yu¨ksel and Merve O¨calan Abstract The epithelial barrier is the basic unit that ensures the continuation of life for all living things. It provides separation of living cells or organelles from nature and microenvironment. Thus, life and functions continue. It is the same for the human organism. However, the normal properties of this epithelial barrier may differ in each organ and tissue. The two most important barriers that separate humans from nature and their microenvironment are the respiratory tract and the gastrointestinal system. The respiratory tract continues from the tip of the nose to the alveola. The epithelial barrier in the respiratory tract has to be semipermeable in places. However, the increase in permeability exceeding the limit is the cause of the diseases and the increase in clinical weight. Therefore, measuring the level of epithelial permeability in these units is important for understanding experimental models, disease cause, clinical severity, and prognosis. In this article, the measurement of epithelial permeability in the respiratory tract will be discussed with in vitro, in vivo aspects and methods. Key words Airway, Epithelial permeability, Measurement, Method, Protocol

1

Introduction Human airway starts from nasal orifice and ends in the alveoli. The entire airway is covered with epithelial cells. Airway epithelial cells are an important part of the innate immune system. Epithelial cells produce anti-microbial peptides, chemokines, and cytokines that recruit and activate other cell types and promote pathogen clearance and establish mucociliary clearance. [1] Airway epithelial cells also form a barrier against the outside world, as in the skin and gastrointestinal tract. This barrier comprised of airway surface liquids, mucus, and apical junctional complexes (AJC) that form between neighboring cells [2]. AJC, which has an important role in epithelial defense, consist of tight junctions (TJs), located at the most apical and the underlying adherens junctions (AJs). (Fig. 1, [3]) Epithelial tight junctions and adherens junctions establish cellcell contact, cell polarity, and also regulate the paracellular movement of ions and macromolecules [2]. Especially TJs regulate paracellular transport of ions and some molecules while AJs are responsible for cell-cell adhesion.

149

150

Hasan Yu¨ksel and Merve O¨calan

Fig. 1 Cell to cell epithelial barrier components of airway barrier

Intercellular junctions were first identified using a light microscope. Afterwards, with the advent of electron microscopy (EM), especially freeze-fracture techniques, TJ was observed to have linear strands just below the apical membrane and proceed as parallel bands. The first systematic analysis of airway epithelial tight junction morphology by EM was reported in 1992 [4]. Next, Jeffery et al. compared the airway junction morphology in the lungs in patients with and without cystic fibrosis (CF). They found some differences in merger complexity especially disorganized strands that sometimes extended below the typical apical belt [5]. Tight junctions constitute the most important part of the physical barrier function of epithelium [6, 7]. TJs establish the

Measurement of Airway Epithelial Permeability: Methods and Protocols

151

major barrier to paracellular ion and fluid traffic between the inner and outer environments [8]. TJ proteins contain 3 major transmembrane proteins: (1) members of claudin family; (2) tight junction-associated MARVEL protein family members: occludin, tricellulin, and MARVELD3; and (3) immunoglobulin-like proteins, such as junctional adhesion molecule (JAM) and coxsackie adenovirus receptor (CAR) [9]. Epithelial AJ (adherens junction) proteins are mainly composed of E-cadherin and members of nectin family proteins [10]. E-cadherin is a calcium-dependent adhesion molecule expressed on epithelial cells and essentially mediates homophilic cell-cell adhesion. E-cadherin is believed to be necessary for the maintenance of cell structure and other TJ structures. When E-cadherin is not properly expressed in the epidermis, TJ proteins ZO-1, occludin, and claudin are delocalized and TJs become distorted [11]. Peripheral membrane proteins form the cytosolic plaque of TJ and AJ. They cluster and stabilize adhesive components of the AJC. Especially zonula occludens (ZO) proteins bind the intracellular domains of TJ components with actin binding proteins such as cortactin. Proteins such as afadin, cingulin, β-catenin, and pI20 catenin are also other components of the cytosolic plaque that connects the cytoskeleton with TJ and E-cadherin [3]. On the other hand, polarity proteins is important for assembly of TJ and AJ properly. Different families of proteins have been identified including the PAR complex (containing six par genes and protein kinase C3), the Crumbs complex (composed of Crumbs, PALS1, and PATJ), and the Scribble complex (containing scribble, Dlg, and Lgl) as polarity proteins. Barrier function is usually studied by growing epithelial cells in vitro on semipermeable membranes. A current concern in this model is to understand how findings from monolayer epithelium extrapolate to multicellular epithelium under real-world conditions. This concern can be alleviated by the use of primary epithelial cells differentiating at the air-liquid interface (ALI) that are very similar to in situ epithelial cells [12]. In order to demonstrate the epithelial barrier permeability, in vivo animal experiments and in vitro cell cultures as also organoid models are studied most recently. Established methods of barrier function analyses from two-dimensional cultures have to be adjusted to the analysis of three-dimensional organoid structures. Two general approaches are used to study junction function: (1) permeability to small ions can be measured by analyzing transepithelial electrical resistance (TEER) and (2) permeability to probes of different sizes and shapes can be estimated by measuring flux across the monolayer. This is typically expressed normalized to surface area and concentration as apparent permeability: Papp ¼ F/ (S  A0), where F ¼ flow rate, S ¼ surface area, and A0 ¼ starting concentration [13].

152

2

Hasan Yu¨ksel and Merve O¨calan

Materials Changes in AJC structure and lack or overexpression of AJC proteins alter epithelial barrier function and increase epithelial permeability. Some claudins that make up the structure of AJC increase paracellular permeability (“pore forming” or “leaky” claudins, for example claudin-2). Some claudins interact in a homo- or heterotypic manner to improve barrier integrity (e.g., klaudin-1). In previous some studies, it has been shown that overexpression of occludin in MDCK cells increases epithelial permeability with 3- or 4-kD dextran [14–16]. The occurrence of mucosal inflammation in the airway increases the epithelial permeability. Some environmental exposures have been shown to enhance airway epithelial permeability without inducing cytotoxicity. These include air pollution components especially cigarette smoke, respiratory viruses, and allergens. In many previous studies, air pollution components such as ozone, high concentrations of diesel exhaust particles, and carbon nanotubes were shown to reduce TEER and/or to increase permeability in different lung epithelial cell lines [11, 17–19]. Many studies have shown that exposure to cigarette smoke (CS) disrupts airway epithelial integrity, decreasing TEER and increasing permeability [20, 21]. Also, levels of E-cadherin and β-catenin were found to be low in the epithelium after CS exposure. This situation is also associated with epithelial mesenchymal transition. Infections with some viruses such as influenza lead to barrier dysfunction by causing epithelial cell death due to direct cytopathic effect or indirectly via immune cell cytotoxicity. Some respiratory viruses such as Coxsackie B, adenoviruses, and reoviruses also cause apical junction disruption without inhibiting cell viability. In studies on this subject, it has been shown that junctional components can serve as receptors for viruses and promote cell entry [22–24]. Cell culture studies with 16HBE (human epithelial bronchial cells) and PBEC (primary airway epithelial cells) have shown that infections with RV and RSV resulted in sustained decreases in TEER and increased paracellular permeability [25–28]. Recent studies have documented defects in airway barrier structure and the presence of dysfunctional epithelial AJC in the asthmatic patients. However, the mechanisms and consequences of airway inflammation are not clear. Inhaled allergens, pollution particles, and respiratory viruses can disrupt barrier integrity, which may represent a risk factor for allergen sensitization. Some inflammatory cytokines can also cause barrier dysfunction in the asthmatic airway. In a study by De Boer et al., a decrease in alphacatenin, E-cadherin, and ZO-1 expression was observed in bronchial biopsies taken from patients with asthma [29]. In a research by Xiao et al., the expression of ZO-1 was significantly reduced in epithelial biopsies. Second, TEER was

Measurement of Airway Epithelial Permeability: Methods and Protocols

153

significantly reduced in epithelial cells after 21 days of culture in vitro and correlated with asthma severity. Third, TEER decreased in airway epithelial cells from asthmatic subjects and increased permeability to 4- and 20-kD dextrans [30].

3

Methods Although our knowledges about epithelial barrier increase day by day, there is not only single model that can fully explain how changes in the epithelial barrier and AJC structure affect paracellular permeability. When looking at the studies on this subject, the following thought comes to the fore. I˙ntercellular junctions are not static resistors but rather dynamic structures with discrete probabilities of being open or closed [31, 32]. In this study, we will summarize the methods used to measure the epithelial barrier permeability until today.

3.1 Transepithelial/ Transendothelial Electrical Resistance (TEER)

Transepithelial/transendothelial electrical resistance (TEER) is a quantitative technique used to measure tight junction integrity in cell culture models of endothelial and epithelial monolayers. TEER measurements are based on measuring ohmic resistance or impedance over a wide range of frequencies. Measurement of TEER and other flux molecules are both indicators of the integrity of the tight junctions and of the cell monolayer. TEER reflects the ionic conductance of the paracellular pathway in the epithelial monolayer, whereas the flux of non-electrolyte tracers indicates the paracellular water flow, as well as the pore size of the tight junctions [33]. The advantages of the TEER method are that it is non-invasive and suitable for monitoring living cells at various growth and differentiation stages. The disadvantages are that different measurements may occur depending on factors such as temperature, media formulation, and cell number [34].

3.2 TEER Measurement Methods

The electrical resistance of a cellular monolayer is measured in ohms. It is a quantitative measure of the barrier integrity [35]. It contains a semipermeable filter that consists of a cellular monolayer cultured on. This filter defines a partition for apical (or upper) and basolateral (or lower) compartments (Fig. 2, [34]). For electrical measurements, two electrodes are used, with one electrode placed in the upper compartment and the other in the lower compartment, and the electrodes are separated by the cellular monolayer. In theory, the ohmic resistance can be determined by applying a direct current (DC) voltage to the electrodes and measuring the resulting current. The ohmic resistance is calculated based on Ohm’s law as the ratio of the voltage and current. DC currents can damage both the cells and the electrodes. Therefore, an alternating current (AC) voltage signal with a square waveform is

3.2.1 Ohm’s Law Method

154

Hasan Yu¨ksel and Merve O¨calan

Fig. 2 TEER measurement with chopstick electrodes. The total electrical resistance includes the ohmic resistance of the cell layer RTEER, the cell culture medium RM, the semipermeable membrane insert RI, and the electrode medium interface REMI

applied. In the widely used and a commercial TEER measurement system known as an Epithelial Voltohmmeter (EVOM; World Precision Instruments, Sarasota, FL) [36], an AC square wave at a frequency of 12.5 Hz is used to avoid any charging effects on the electrodes and the cell layer. The EVOM system uses a pair of electrodes known as a STX2/“chopstick” electrode pair. Each stick of the electrode pair contains a silver/silver chloride pellet for measuring voltage and a silver electrode for passing current. Firstly, the blank resistance (RBLANK) of the semipermeable membrane only (without cells) is measured. Then the resistance across the cell layer on the semipermeable membrane (RTOTAL) is measured. The cell-specific resistance (RTISSUE), in units of Ω, can be obtained as: RTISSUE ðΩÞ ¼ RTOTAL  RBLANK where resistance is inversely proportional to the effective area of the semipermeable membrane (MAREA ), which is reported in units of cm2 [37]. RTISSUE ðΩÞ α 1=M AREA ðcm2 Þ TEER values are reported (TEERREPORTED) in units of Ω.cm2 and calculated as: TEERREPORTED ¼ RTISSUE(Ω)  MAREA(cm2) [37, 38]

Measurement of Airway Epithelial Permeability: Methods and Protocols

155

Fig. 3 TEER measurement concept based on impedance spectroscopy

3.2.2 Impedance Spectroscopy

Impedance spectroscopy demonstrates TEER values better than DC/single frequency AC measurement systems [39]. It can give information about the capacitance of the cell layer. Impedance spectroscopy is performed by applying a small amplitude AC excitation signal with a frequency sweep and measuring the amplitude and phase response of the resulting current (Fig. 3, [34]). An automated measurement system (cellZscope, nanoAnalytics GmbH, Germany) has been developed for measuring the transendothelial/epithelial impedance of various barrier-forming cells cultures [34]. An equivalent circuit analysis of the measured impedance spectrum is performed to obtain the electrical parameters that can be applied to characterize the cellular barrier properties (Fig. 4a, adapted from Benson et al. [35]). In this circuit, the current can flow through the junctions between cells (paracellular route) or through the cell membrane of the cells (transcellular route). The tight junction proteins in the paracellular route contribute to an ohmic resistance (RTEER) in the equivalent circuit. Each lipid bilayer in the transcellular route contributes to a parallel circuit [35] consisting of ohmic resistance (Rmembrane) and an electrical capacitance (CC). The high values of Rmembrane cause the current to mostly flow across the capacitor and allow Rmembrane to be ignored (Fig. 4b, [35]). The impedance spectrum observed will have a nonlinear frequency dependency as shown in Fig. 4c [35]. There are three distinct frequency regions in the impedance spectrum and some circuit elements dominate this impedance. In the low-frequency range, the impedance signal is dominated by CE (capacitance of the measurement electrodes). In the midfrequency range, the impedance signal is dominated by circuit elements related to the cells, namely, RTEER and CC. In the high-frequency range, CC and CE provide a more conductive path and the impedance signal is dominated by Rmedium. These equivalent circuit parameters can be estimated by fitting the experimental impedance spectrum data

156

Hasan Yu¨ksel and Merve O¨calan

Fig. 4 (a) A typical equivalent circuit diagram that can be applied to analyze the impedance spectrum of cellular systems. (b) Simplified equivalent circuit. (c) A typical impedance spectrum with distinct frequencydependent regions. (Adapted from Benson et al.)

to the equivalent circuit model using nonlinear least-squares fitting techniques to obtain the best fit parameters [34]. 3.2.3 Other Systems Available for TEER Measurements Ussing Chamber

The Ussing chamber was an apparatus developed by the Danish biologist Hans H. Ussing [40] in the early 1950s. It was developed to measure the active transport of sodium in frog skin epithelium. Ussing and Zerahn demonstrated that the rate of active transport of ions can be calculated as an electric current across the skin if both sides are at the same potential and have similar solutions [41]. Thereby, there is no net transfer of passive ions and the electric current (Isc) is only caused by active transport processes. With Ussing chamber system, Vte (transepithelial voltage), TEER, and Isc are measured. A primary advantage of this system is the high accuracy of the measurement because of the uniform current distribution along the epithelial sheet [42].

Measurement of Airway Epithelial Permeability: Methods and Protocols

157

Transwells

Transwell permeable supports are cell culture to be used with standard well plates. There is an insert body with a bottom filter plate. Membrane filter acts as a cell growth substrate and permits particles to diffuse through the filter pores. The advantage of this method is that there is a basal side where cells on permeable supports can take up molecules and secrete them. This provides a better imitation of in vivo. Moreover, Transwell system facilitates the collection of samples from both tissue sides and therefore permits transport studies. Electrophysiological measurement of the epithelial barrier is provided with chopstick-like tetrapolar electrodes. A disadvantage of chopstick-like systems is the proper positioning of the sticks and the non-uniform current distribution when large membrane filters are used (>12 mm in diameter) [43]. To correct this limitation, stick electrodes can be replaced with so-called “Endohm series” (World Precision Instruments, Sarasota, FL, USA) [42].

ECIS (Electric CellSubstrate Impedance Sensing)

ECIS is a method by which cells are cultured on a surface containing sensing electrodes (Fig. 5d [44]). ECIS contains a small circular gold electrode of 250 μm in diameter surrounded by a large

Fig. 5 Transepithelial electrical measurement techniques. (a) Original diagram representation of the Ussing chamber in 1951. (b) Detailed parts of the Ussing chamber. (c) Schematic representation of chopstick-like electrodes for use with standard Transwell inserts. (d) Electric cell-substrate impedance sensing system

158

Hasan Yu¨ksel and Merve O¨calan

electrode. Advantage of the ECIS technology is more sensitive to cell motility and shape of small populations of cells. And it can be repeatable due to the fixed electrodes. As a disadvantage, measurements with ECIS electrodes are only representative of small areas. In addition, transport studies are not allowed in ECIS cultureware since there is no basal compartment. 3.3 Two-Path Impedance Spectroscopy

Solutes are transported across epithelial cell layers via transcellular and paracellular pathways. The transcellular pathway leads across the apical and basolateral cell membrane, whereas the paracellular pathway is directed through the tight junction. Two-path impedance spectroscopy method is based on a transepithelial impedance measurement in specialized Ussing chambers, combined with a Ca2+-dependent modulation of Rpara through EGTA and flux measurements of fluorescein (as a paracellular marker) [45]. As it discriminates between the trans- and paracellular pathway, it is referred to as two-path impedance spectroscopy. If transepithelial flux of a truly paracellular marker is measured, this flux should be directly proportional to the paracellular conductance, Gpara (¼1/Rpara): G epi ¼ G trans þ s  Flux G epi ¼ G trans þ G para ¼ G trans þ s  Flux Gepi ¼ 1/Repi (¼1/R,t  R,alt) ve Gtrans ¼ 1/R,trans (Fig. 6b) ¼ 1/(Rap + Rbl) (Fig. 6c) [45]. (Abbreviations: Flux:fluorescein flux, s:contant, C:capacitance, R: resistance, G: conductance(¼1/R), Z:impedance, t:total, sub: subepithelial, epi:epithelial, trans:transcellular, para:paracellular, ap:apical, bl:basolateral.) MDCK (Madin–Darby Canine Kidney cell) and HT-29/B6 (colonic epithelium cell) cell layers were used in this method. New studies are needed in airway epithelial cells.

3.4 FITC (Fluorescein Isothiocyanate)

Fluorescein isothiocyanate is a water soluble, photostable, biocompatible molecule. The stable and high intense fluorescence emission allows easy detection of small particles [46]. Fluorescent labeled polymers are used in epithelial permeability studies by binding with molecules such as dextran or electrolytes such as sodium. FITCdextran flux measurements is usually used as a marker of epithelial layer integrity. FITC-dextran is added to cell cultures apically at given time points. After a certain period after addition, the FITC intensity of basolateral fluids is measured using a spectrophotometer at a specific wavelength (e.g., an excitation wavelength of 492 nm and emission wavelength of 520 nm). In a research with RSV infected 16HBE14o- human bronchial epithelial cells, epithelial permeability was performed by measuring

Measurement of Airway Epithelial Permeability: Methods and Protocols

159

Fig. 6 One- and two-path impedance spectroscopy models. (a) Equivalent electrical circuit for one path impedance measurements, that only discriminate between subepithelial (Rsub) and epithelial (Repi) resistance, and the epithelial capacity Cepi. (b) Equivalent electrical circuit for two-path impedance measurements used in this study. Here, the epithelial resistance Repi is composed of a paracellular (Rpara) and a parallel transcellular (Rtrans) resistance. Under DC conditions or AC conditions at very low frequencies Repi ¼ Rpara Rtrans/ (Rpara + Rtrans). (c) Equivalent electrical circuit for a model discriminating between the apical (index ap) and the basolateral (index bl) side of an epithelium. In this model, the capacitance Cepi of the models presented in A and B is composed of Cap and Cbl (Cepi ¼ Cap  Cbl/(Cap + Cbl). Under DC conditions or AC conditions at very low frequencies Repi ¼ Rpara  (Rap + Rbl)/ (Rpara + Rap + Rbl)

the flux of apically added fluorescein isothiocyanate (FITC)conjugated 3-kDa dextran. RSV infection decreased resistance and increased the permeability of polarized airway epithelial cells [47]. In a study conducted by Soyka et al., it was shown that epithelial integrity was disrupted by decreased expression of TJ proteins under the effect of IFN GAMA and IL-4 in patients with chronic rhinosinusitis and epithelial permeability increased with FITC-dextran [48]. 3.5

Mannitol

Mannitol is one of the molecules used to show epithelial permeability. It is administered at a certain concentration from the apical part of the cell line. Flux rate is evaluated by comparing samples collected from the basolateral area. In a previous study [49] with human bronchial epithelial cell culture (16HBE14o-) model, the basal-lateral medium was aspirated and replaced with 15 ml of medium containing 0.1 mM, 0.1 μCi/ml 14C-d-mannitol and incubated at 37  C. Triplicate 50-μl samples were taken from the basolateral medium to determine the specific activity via liquid scintillation counting (LSC). Duplicate 250-μl samples were taken from the apical side at either 60 or 90 min for LSC to determine flux rates (14C-d-mannitol flux rate, Jm). The flux rate (in picomoles/min/cm2) was calculated for 14C-d-mannitol diffusing across the cell layer.

160

Hasan Yu¨ksel and Merve O¨calan

3.6

Albumin

Albumin is one of the molecules used in studies to show airway epithelial barrier permeability. In a study by Eutamene et al. [50], showed that LPS due to P. aeruginosa increased airway epithelial permeability as both in vivo and in vitro method. Evaluation of the permeability of the airway epithelial barrier (AEB) was evaluated by measuring the accumulation of residual [125I] albumin, the airspace protein tracer, and [125I] albumin in plasma at in vivo model. In a research conducted on HBECs obtained from asthmatic and non-asthmatic individuals, the effect of air, O3, and NO2 on epithelial permeability was investigated [18]. 14C-bovine serum albumin (14C-BSA) was used in this study. Just before exposure, cells were incubated for 30 min in the presence of 0.025 μCi 14C BSA. At the end of this incubation, the medium in each insert well was collected and analyzed for total radioactivity, by liquid scintillation counting in a Beckman LS6500 scintillation counter.

3.7

Lucifer Yellow

The permeability of the epithelial cells can also be studied by measuring the passive transport of small hydrophilic molecules such as lucifer yellow across the monolayer. This molecule mainly passes the monolayer via the paracellular route, i.e., through the tight junctions (TJs), and can be used to get information about the leakiness of the TJs. In these studies, 4.5 μM LY solution is usually given from the apical side and samples are collected from the basolateral side at certain time intervals. As well as in monolayer samples, this method can be studied in 2-dimensional cultures and new 3D organoids. In a study with intestinal epithelial cell organoid cultures, Lucifer yellow was used to demonstrate epithelial permeability. Upon beginning 1 mM lucifer yellow was added to the medium, organoids were imaged for 70 min by confocal microscopy with an interval of 5 min. Densitometric analysis was performed (using Leica LAS X Core software). For each time point, lucifer yellow was quantified as fluorescence. Fluorescence intensity inside the organoid was normalized to the mean total fluorescence intensity [51].

4

Notes In order to obtain accurate epithelial barrier permeability measurements, some variables should be optimized methodologically. To take a look at some of them, in cell cultures, number of cell layers and passage number affect the TEER measurements. This should be taken into account and passage number should be recorded. The origin of cell lines and the different solutions used to maintain these cells in culture can affect the process of spontaneous differentiation. In a study comparing a commercial mixture, MITO+ serum

Measurement of Airway Epithelial Permeability: Methods and Protocols

161

extender, and a fetal bovine serum symmetric medium (complete Dulbecco’s modified Eagle’s medium supplemented with 10% FBS) for the Caco-2 cells, it was resulted with significantly lower TEER values in MITO+ serum extender [52]. On the other hand, the culture period is considered to be important for the formation of a tight junction in cell lines. Cell cultures should be observed in optimum time after seeding. TEER is an epithelial permeability measurement method that the passage of small ions is measured. In the TEER methods, some factors may affect the TEER values such as temperature, media formulation, and cell number [34]. Therefore, TEER measurements should be conducted in an incubator at 37  C. If TEER measurements are performed at room temperature, cell physiology and function could be affected by any temperature fluctuation and results may change. A mathematical method has been developed to correct TEER values at temperature recorded. This is referred to as temperature-corrected TEER (tcTEER) [53]. This application minimizes the temperature fluctuations that the cells are exposed to that could be detrimental to their function.

5

Conclusion Disruption or dysfunction of the epithelial barrier in the airway is the most important cause of congenital airway diseases and the most important result of acquired airway diseases. Disease development is also often associated with this in experimental models. Experimental and clinical determination of the degree of my illness and determination of recovery are related to the state of the airway epithelium barrier function. The strength of the barrier is also related to its permeability state. That is, evaluating the functional state of the barrier permeability and its dysfunction will give the most important result. Therefore, it is very important to use and develop methods and protocols to evaluate airway barrier permeability.

References 1. Schleimer RP, Kato A, Kern R, Kuperman D, Avila PC (2007) Epithelium: at the interface of innate and adaptive immune responses. J Allergy Clin Immunol 120:1279–1284 2. Georas SN, Rezaee F (2014) Epithelial barrier function: at the front line of asthma immunology and allergic airway inflammation. J Allergy Clin Immunol 134(3):509–520. https://doi. org/10.1016/j.jaci.2014.05.049 3. Rezaee F, Georas SN (2014) Breaking barriers. New insights into airway epithelial barrier

function in health and disease. Am J Respir Cell Mol Biol 50(5):857–869 4. Godfrey RW, Severs NJ, Jeffery PK (1992) Freeze-fracture morphology and quantification of human bronchial epithelial tight junctions. Am J Respir Cell Mol Biol 6:453–458 5. Godfrey RW, Severs NJ, Jeffery PK (1993) Structural alterations of airway epithelial tight junctions in cystic fibrosis: comparison of transplant and postmortem tissue. Am J Respir Cell Mol Biol 9:148–156

162

Hasan Yu¨ksel and Merve O¨calan

6. Swindle EJ, Collins JE, Davies DE (2009) Breakdown in epithelial barrier function in patients with asthma: identification of novel therapeutic approaches. J Allergy Clin Immunol 124:23–34 7. Kojima T, Bas¸a M, Takano K, Kuros M, Ohkuni T et al (2013) Regulation of tight junctions in upper airway epithelium. Biomed Res Int 2013:947072 8. Yuksel H, Turkeli A, Taneli F, Horasan GD, Toprak Kanik E et al (2014) E-cadherin as an epithelial barrier protein in exhaled breath condensate. J Breath Res 8(4):046006 9. Schulzke JD, Fromm M (2009) Tight junctions: molecular structure meets function. Ann N Y Acad Sci 1165:1–6 10. Niessen CM, Gottardi CJ (2008) Molecular components of the adherens junction. Biochim Biophys Acta 1778(3):562–571 11. Nawijn MC, Hackett TL, Postma DS, van Oosterhout AJ, Heijink IH (2011) E-cadherin: gatekeeper of airway mucosa and allergic sensitization. Trends Immunol 32:248–255 12. You Y, Richer EJ, Huang T, Brody SL (2002) Growth and differentiation of mouse tracheal epithelial cells: selection of a proliferative population. Am J Physiol Lung Cell Mol Physiol 283:L1315–L1321 13. Strengert M, Knaus UG (2011) Analysis of epithelial barrier integrity in polarized lung epithelial cells. Methods Mol Biol 763:195–206 14. Balda MS, Whitney JA, Flores C, Gonza´lez S, Cereijido M, Matter K (1996) Functional dissociation of paracellular permeability and transepithelial electrical resistance and disruption of the apical-basolateral intramembrane diffusion barrier by expression of a mutant tight junction membrane protein. J Cell Biol 134:1031–1049 15. McCarthy KM, Francis SA, McCormack JM, Lai J, Rogers RA et al (2000) Inducible expression of claudin-1-myc but not occludin-VSV-G results in aberrant tight junction strand formation in MDCK cells. J Cell Sci 113:3387–3398 16. Van Itallie CM, Fanning AS, Holmes J, Anderson JM (2010) Occludin is required for cytokine-induced regulation of tight junction barriers. J Cell Sci 123:2844–2852 17. Yu XY, Takahashi N, Croxton TL, Spannhake EW (1994) Modulation of bronchial epithelial cell barrier function by in vitro ozone exposure. Environ Health Perspect 102:1068–1072 18. Bayram H, Rusznak C, Khair OA, Sapsford RJ, Abdelaziz MM (2002) Effect of ozone and nitrogen dioxide on the permeability of bronchial epithelial cell cultures of non-asthmatic and asthmatic subjects. Clin Exp Allergy 32:1285–1292

19. Lehmann AD, Blank F, Baum O, Gehr P, Rothen-Rutishauser BM (2009) Diesel exhaust particles modulate the tight junction protein occludin in lung cells in vitro. Part Fibre Toxicol 6:26 20. Schamberger AC, Mise N, Jia J, Genoyer E, Yildirim AO et al (2014) Cigarette smokeinduced disruption of bronchial epithelial tight junctions is prevented by transforming growth factor-beta. Am J Respir Cell Mol Biol 50(6):1040–1052 21. Nishida K, Brune KA, Putcha N, Mandke P, O’Neal WK et al (2017) Cigarette smoke disrupts monolayer integrity by altering epithelial cell-cell adhesion and cortical tension. Am J Physiol Lung Cell Mol Physiol 313(3): L581–L591 22. Bergelson JM, Cunningham JA, Droguett G, Kurt-Jones EA, Krithivas A et al (1997) Isolation of a common receptor for Coxsackie B viruses and adenoviruses 2 and 5. Science 275:1320–1323 23. Barton ES, Forrest JC, Connolly JL, Chappell JD, Liu Y et al (2001) Junction adhesion molecule is a receptor for reovirus. Cell 104:441–451 24. Coyne CB, Bergelson JM (2006) Virusinduced Abl and Fyn kinase signals permit coxsackievirus entry through epithelial tight junctions. Cell 124:119–131 25. Sajjan U, Wang Q, Zhao Y, Gruenert DC, Hershenson MB (2008) Rhinovirus disrupts the barrier function of polarized airway epithelial cells. Am J Respir Crit Care Med 178:1271–1281 26. Comstock AT, Ganesan S, Chattoraj A, Faris AN, Margolis BL et al (2011) Rhinovirusinduced barrier dysfunction in polarized airway epithelial cells is mediated by NADPH oxidase 1. J Virol 85:6795–6808 27. Singh D, McCann KL, Imani F (2007) MAPK and heat shock protein 27 activation are associated with respiratory syncytial virus induction of human bronchial epithelial monolayer disruption. Am J Physiol Lung Cell Mol Physiol 293:L436–L445 28. Rezaee F, DeSando SA, Ivanov AI, Chapman TJ, Knowlden SA et al (2013) Sustained protein kinase D activation mediates respiratory syncytial virus-induced airway barrier disruption. J Virol 87:11088–11095 29. De Boer WI, Sharma HS, Baelemans SM, Hoogsteden HC, Lambrecht BN, Braunstahl GJ (2008) Altered expression of epithelial junctional proteins in atopic asthma: possible role in inflammation. Can J Physiol Pharmacol 86:105–112

Measurement of Airway Epithelial Permeability: Methods and Protocols 30. Xiao C, Puddicombe SM, Field S, Haywood J, Broughton-Head V et al (2011) Defective epithelial barrier function in asthma. J Allergy Clin Immunol 128:549–556, e12 31. Shen L, Weber CR, Raleigh DR, Yu D, Turner JR (2011) Tight junction pore and leak pathways: a dynamic duo. Annu Rev Physiol 73:283–309 32. Schneeberger EE, Lynch RD (2004) The tight junction: a multifunctional complex. Am J Physiol Cell Physiol 286:C1213–C1228 33. Zucco F, Batto AF, Bises G (2005) An interlaboratory study to evaluate the effects of medium composition on the differentiation and barrier function of Caco-2 cell lines. Altern Lab Anim 33:603–618 34. Srinivasan B, Kolli AR, Esch MB, Abacı HE, Shuler ML, Hickman JJ (2015) TEER measurement techniques for in vitro barrier model systems. J Lab Autom 20(2):107–126 35. Benson K, Cramer S, Galla HJ (2013) Impedance-based cell monitoring: barrier properties and beyond. Fluids Barriers CNS 10:5 36. Epithelial Voltohmmeter (2014) http://www. wpi-europe.com/products/cell-and-tissue/ teer-measurement/evom2.aspx. Accessed 13 Nov 2014 37. Haorah J, Schall K, Ramirez SH (2008) Activation of protein tyrosine kinases and matrix metalloproteinases causes blood-brain barrier injury: novel mechanism for Neurodegeneration associated with alcohol abuse. Glia 56:78–88 38. Watson PMD, Paterson JC, Thom G (2013) Modelling the endothelial blood-CNS barriers: a method for the production of robust in vitro models of the rat blood-brain barrier and blood-spinal cord barrier. BMC Neurosci 2013:14 39. Douville NJ, Tung YC, Li R (2010) Fabrication of two-Layered Channel system with embedded electrodes to measure resistance across epithelial and endothelial barriers. Anal Chem 82:2505–2511 40. Larsen EH (2009) Hans Henriksen Ussing. 30 December 1911 – 22 December 2000. Biogr Mem Fellows R Soc 55:305–335 41. Ussing HH, Zerahn K (1951) Active transport of sodium as the source of electric current in the short-circuited isolated frog skin. Acta Physiol Scand 23:110–127 42. Yeste J, Illa X, Alvarez M, Villa R (2018) Engineering and monitoring cellular barrier models. J Biol Eng 12:18

163

43. Yeste J, Illa X, Gutie´rrez C, Sole´ M, Guimera` A, Villa R (2016) Geometric correction factor for transepithelial electrical resistance measurements in transwell and microfluidic cell cultures. J Phys Appl Phys 375401:49 44. Giaever I, Keese CR (1986) Use of electric fields to monitor the dynamical aspect of cell behavior in tissue culture. IEEE Trans Biomed Eng 33:242–247 45. Krug SM, Fromm M, Gu¨nzel D (2009) Two-path impedance spectroscopy for measuring paracellular and transcellular epithelial resistance. Biophys J 97(8):220211 46. Caprifico AE, Polycarpou E, Foot PJS, Calabrese G (2020) Biomedical and pharmacological uses of fluorescein isothiocyanate chitosanbased nanocarriers. Macromol Biosci e2000312 47. Rezaee F, DeSando SA, Ivanov AI, Chapman TJ, Knowlden SA et al (2013) Sustained protein kinase D activation mediates respiratory syncytial virus-induced airway barrier disruption. J Virol 87(20):11088–11095 48. Soyka MB, Wawrzyniak P, Eiwegger T, Holzmann D, Treis A et al (2012) Defective epithelial barrier in chronic rhinosinusitis: the regulation of tight junctions by IFN-g and IL-4. J Allergy Clin Immunol 130 (5):1087–1096 49. Callaghan PJ, Ferrick B, Rybakovsky E, Thomas S, Mullin JM (2020) Epithelial barrier function properties of the 16HBE14o- human bronchial epithelial cell culture model. Biosci Rep 40: BSR20201532 50. Eutamene H, Theodorou V, Schmidlin F, Tondereau V, Garcia-Villar R et al (2005) LPS-induced lung inflammation is linked to increased epithelial permeability: role of MLCK. Eur Respir J 25:789–796 51. Bardenbachera M, Ruderb B, BritzenLaurenta N, Schmide B, Waldnerb M et al (2019) Permeability analyses and three dimensional imaging of interferon gamma- induced barrier disintegration in intestinal organoids. Stem Cell Res 35:101383 52. Ferruzza S, Rossi C, Sambuy Y (2013) Serumreduced and serum-free Media for Differentiation of Caco-2 cells. ALTEX 30:159–168 53. Blume LF, Denker M, Gieseler F (2010) Temperature corrected transepithelial electrical resistance (TEER) measurement to quantify rapid changes in paracellular permeability. Pharmazie 65:19–24

Methods in Molecular Biology (2021) 2367: 165–175 DOI 10.1007/7651_2020_310 © Springer Science+Business Media New York 2020 Published online: 18 August 2020

Vascular Permeability Assays In Vivo Mir S. Adil and Payaningal R. Somanath Abstract Whereas physiological vascular permeability (VP) mediates selective transport of plasma, electrolytes, proteins, and cells across an intact endothelial barrier, pathological VP results in the loss of endothelial barrier integrity. Whereas physiological VP is a feature of regular host defense and tissue repair, compromised barrier function may lead to aberrant vascular leakage, concurrent tissue edema, and inflammation eventually causing life-threatening conditions such as acute lung injury or acute respiratory distress syndrome, cancer, kidney injury, etc. Measurement of VP helps to identify, design, and optimize anti-leak therapies. Further, it can define the effect of a stimulus or a gene modulation in endothelial-barrier regulation. The degree of VP can be of importance to determine the stage of cancer and disease prognosis. This chapter discusses Miles assay, which is a well-established, relatively simple, and a reliable in vivo technique to assess VP as a surrogate measurement. Although a reliable technique, Miles assay is timeconsuming, and the technique does not consider the compounding factors that may increase VP independently of endothelial-barrier regulation, such as blood pressure or blood flow. As an alternative, we describe fluorescein isothiocyanate-dextran lung permeability assay, a method that can also be adapted to measure VP and edema in other organs such as the brain and kidney. Keywords Evans blue, FITC dextran, In vivo permeability, Miles assay, Vascular permeability

1

Introduction Vascular permeability (VP) is a characteristic of the capillary wall to prevent movement of plasma or cells guided by a physical force [1]. It is a highly complex and coordinated event primarily important in cardiovascular diseases, cancer, and inflammation [2]. Endothelial cells line the lumen side of the blood vessel and form a selectively permeable barrier that helps in transport between blood and the interstitial space of all organs. The development and maintenance of these barriers are crucial for organ growth and performance [3]. VP also plays an integral role in the regulation of leukocyte infiltration and leakage of plasma elements to sustain tissue homeostasis. In vivo, it is thought to be controlled predominantly by blood flow and endothelial-barrier function [4].

165

166

Mir S. Adil and Payaningal R. Somanath

1.1 Mechanism of Vascular Permeability

Growth factors, cytokines, and other stress-related molecules regulate an adequate VP barrier through tight cell-to-cell junctions. Disruption of these barriers may result in increased permeability and uncontrolled vascular leakage [3]. Whereas increased VP may result from a physiological feature of regular host defense needed for tissue repair and wound healing, it can also be destructive by providing simple access for tumor cell invasion to the bloodstream [5]. Under pathological situations, the compromised endothelialbarrier function may lead to vascular leakage, concurrent tissue edema, and inflammation [5] eventually causing life-threatening conditions, such as cerebral hemorrhage and severe inflammatory disorders including acute lung injury/acute respiratory distress syndrome, kidney injury, sepsis, as well as cancer [6]. Various inflammatory mediators such as vasoactive amines (histamine, serotonin), vasoactive peptides (substance P, CAP37), complement fragments (C3a, C4a, C5a), and lipid mediators (eicosanoids) affect the function of the vasculature [5]. Several pro- and anti-VP agents such as the vascular endothelial growth factor (VEGF) and angiopoietin1 (Ang-1), respectively, modulate VP [2]. While VEGF is widely accepted as a primary inducer of angiogenesis which begins with vascular leakage [7, 8], Ang-1 tightens endothelial barrier, blocking of which results in disruption of the endothelial barrier [2]. Whereas Src kinases are involved in the opening of the endothelial barrier [9], several studies have demonstrated that Akt1, a serine-threonine kinase [10], is essential for the endothelial-barrier integrity [2], inhibition of which results in vascular leakage [11–13].

1.2 Applications of Vascular Permeability Assessment

l

Knowledge of the mechanism behind vascular permeability during pathological situations will be critical for identifying, designing, and optimizing vascular anti-leak therapies [2, 14, 15].

l

Understanding of VP mechanisms and changes in VP with treatments can be vital for the selection of cancer treatment [13, 15].

l

It is reported to be a critical factor for drug delivery therapies as it affects the convection and consolidation times of drug molecules [15].

l

It can differentiate control animals from animals that have been genetically modified [3].

l

It can define the effect of a stimulus or a modulated gene in terms of inducing changes in endothelial-barrier function [3].

l

The degree of VP can determine the stage of any cancer and hence can be used for the disease prognosis [15].

Vascular Permeability Assays In Vivo l

167

It can also indicate hypoxia in tumors as capillary blood contains unusually high levels of deoxygenated blood, which along with heterogeneous blood flow and large intercapillary distances because of abnormal vasculatures may contribute to microregional hypoxia in tumors [15].

1.3 Miles Assay to Measure Vascular Permeability In Vivo

The Miles assay is a well-established, relatively simple, and commonly employed in vivo technique to assess VP as a surrogate measurement of vascular hyperpermeability [3, 5, 16]. First reported in 1952, the assay has been critical to numerous studies that have discovered mediators of vascular hyperpermeability and their mechanisms of action, such as the VEGF-A that was originally identified as the vascular permeability factor [5, 16].

1.3.1 Working Principle of Miles Assay

The assay is based on the fact that albumin does not cross the endothelial barrier under basal physiological conditions as depicted in Fig. 1 [3]. Evans blue is an azo dye with a high affinity for albumin which is injected in the bloodstream of an experimental animal [3, 17]. Whereas it is expected to be restricted within blood vessels under physiologic conditions, the addition of topical or systemic VP stimulus leads to protein leakage from the blood vessels along with the Evans blue that is bound to albumin [3, 5]. This results in a swift bluish coloration of tissues that have

Fig. 1 Diagrammatic representation of an intact and a leaky blood vessel. Albumin bound to Evans blue dye cannot pass through (a) an intact endothelial barrier, but it can pass through (b) a leaky barrier

168

Mir S. Adil and Payaningal R. Somanath

permeable vessels [3]. The capillary leakage can be quantified by the following [17]: l

Measurement of the mean diameter of the area stained by the leaked dye

l

Assessing the intensity of its color against a set of standards

l

Dye extraction through acid digestion followed by colorimetric measurement

l

Measuring the amount of a radiolabeled or fluorescent marker at the individual injection sites

Successful injection of the dye in the mouse lateral tail vein is critical for the good outcome of the experiment [15]. The increase in optical density values is indicative of the increase in dye retention and hence of vascular leakage [18]. 1.3.2 Limitations of the Miles Assay

1.4 Alternate Method: Fluorescein Isothiocyanate (FITC)Dextran Lung Permeability Assay [21]

l

It is quantified primarily by observation, planimetry, or tissue extraction, which are intrinsically limited by imprecision leading to large variations both within and between animals [5, 17].

l

It is time-consuming, especially if multiple spots are to be assessed [5, 17].

l

The tail-vein injection technique in mice requires extensive practice and skills [3].

l

It is particularly difficult in intradermal studies where tissue extraction is less precise [5].

l

It does not consider the compounding factors that may increase vascular leakage independently of endothelial-barrier regulation, such as blood pressure or blood flow [16].

l

As Evans blue dye relies on its conjugation with albumin to form a large molecule, the limitation is that the smaller unbound molecules of dye, that may escape the clearing process by an osmotic pump, can still diffuse out of intact capillaries even without a significant increase in VP, leading to false-positive conclusions [19].

l

The absorbance of the dye must be corrected for residual heme pigments [20].

An ideal method to assess pulmonary permeability must be simple, feasible, reproducible, and minimally invasive, and the current technique fulfills these criteria to an extent. FITC-dextran is instilled intranasally (i.n.), and the fluorescence intensity (FI) of plasma FITC-dextran is measured and used as the criterion for lung permeability.

Vascular Permeability Assays In Vivo

2

169

Materials

2.1 Miles Assay [2, 3, 14]

Age- and gender-matched C57BL6 and BALB/c mice can be used along with the following: 1. Pipette and tips 2. Spatula 3. Sterile phosphate-buffered saline (PBS) 4. Syringes and needles (27–30 gauge) 5. Mouse restrainer 6. Evans blue dye 7. Dissection instruments 8. Dissection boards and pins 9. Mouse restrainer 10. Biopsy punch 11. Weighing scale 12. Weighing paper 13. Mouse perfusion system 14. Formamide 15. Heat blocks 16. 96-well plate 17. Calorimetric plate reader 18. Digital camera 19. VEGF 20. Ang-1 21. Carbon dioxide chamber 22. Razor blades 23. Trichloroacetic acid 24. Ethanol 25. An osmotic pump

2.2 Fluorescein Isothiocyanate (FITC)Dextran Lung Permeability Assay [21]

1. Age- and gender-matched mice 2. Avertin solution 3. Lipopolysaccharide (LPS), Klebsiella pneumonia, Sigma 4. Sterile PBS 5. FITC-dextran 6. Protein gel loading tips 7. 70% ethanol 8. Dissection instruments

170

Mir S. Adil and Payaningal R. Somanath

9. Dissection boards and pins 10. Syringe and needles (22 gauge) 11. Ethylenediaminetetraacetic acid (EDTA) 12. Centrifuge 13. Synergy H1 plate reader (BioTek) 14. Adenovirus particles expression green fluorescent protein (Ad-GFP) for control and growth factor (Ad-VEGF or Ad-Ang-1) for test

3

Methods

3.1 Miles Assay, e.g., VEGF and Ang-1Mediated VP Assessment [2, 3, 14]

1. Randomly divide the mice into VEGF and Ang-1 groups (see Notes 1 and 2). 2. Anesthetize the mice using isoflurane inhalation (see Note 3) followed by intradermal injection of 30 μl PBS with 20 ng/ml VEGF or 50 ng/ml Ang-1 in the right ear for 30 min. Inject the left ears of mice from both groups with PBS alone as negative controls. 3. Prepare 1% (8 μl/g) Evans blue dye solution in sterile PBS (see Note 4). 4. Aspirate 200 μl Evans blue dye solution into a syringe (see Note 5). 5. Place the mouse into a restraint device so that the animal is not freely mobile and its tail can be handled easily (see Note 6). 6. Hold the tail with the nondominant hand between the thumb and the forefinger (see Note 7). 7. Insert the needle at a 10–15 angle, bevel up, and advance into the lateral tail vein toward the direction of the head (see Note 8). 8. Keep the needle and the syringe parallel to the tail and slowly inject 200 μl of Evans blue solution into the mouse tail vein (see Notes 9 and 10). 9. Repeat steps 4–8 for the rest of the mice from both groups. 10. Place the mice back into the cage for 30 min. 11. Euthanize the mice using a carbon dioxide asphyxiation chamber. 12. Perfuse the mice with sterile PBS using an osmotic pump to remove the excess dye from the vasculature to specifically measure only the dye leaked to the extravascular tissue in the ear (see Note 11). 13. Photograph the ears (see Fig. 2) and cut a circular skin patch from each ear (see Note 12).

Vascular Permeability Assays In Vivo

171

Fig. 2 Ear photographs and calorimetric readings. (a) Representative images (top) of PBS and 30 μl of 20 ng/ml recombinant VEGF administered WT mice and its histogram (bottom) showing calorimetric quantification; (b) representative images (top) of Ad-GFP and Ad-VEGF administered WT mice and its histogram (bottom) showing calorimetric quantification; (c) representative images (top) of PBS and 30 μl of 50 ng/ml recombinant Ang1 administered WT mice and its histogram (bottom) showing calorimetric quantification; (d) representative images (top) of WT mice ears administered with combinations of either Ad-GFP/Ad-VEGF or Ad-VEGF/Ad-Ang1 and its histogram (bottom) showing calorimetric quantification (adapted from ref. [2])

14. Incubate the collected tissue samples in 200 μl formamide for 24 h at 60  C to extract dye. 15. Measure the absorbance of the extracted dye in a colorimetric plate reader (see Note 13). 16. Normalize the absorbance values to their respective tissue weights (see Note 14). 17. To assess the effect of adenovirus particles, inject 30 μl each of Ad-GFP and Ad-VEGF in left and right ears of mice, respectively. 18. Also inject Ad-VEGF + Ad-GFP and Ad-VEGF + Ad-Ang-1 in left and right ears, respectively. 19. To assess the leakage in organs such as lungs, follow the aforesaid steps till step 12 and remove the organs. 20. Rinse, blot dry, weigh, and capture photographs of the isolated organs. 21. Mince the organs with razor blades and homogenize in trichloroacetic acid: ethanol (1:1 v/v). 22. Read absorbance as mentioned in steps 15 and 16. 3.2 Fluorescein Isothiocyanate (FITC)Dextran Lung Permeability Assay [21]

1. Randomly divide the mice into the desired number of groups such as control and acute lung injury (ALI) groups. 2. Anaesthetize mice with appropriate agents (see Note 15).

172

Mir S. Adil and Payaningal R. Somanath

3. Instill 50 μl of PBS with and without LPS intranasally in acute lung injury (ALI) and control groups, respectively (see Notes 16 and 17). 4. After 5 h, introduce 25 μl FITC-dextran in each nostril using protein gel loading tips (see Notes 18 and 19). 5. After instillation, the mouse should be set in a vertical position for 2 min until it breathes steadily. 6. The mice are then sacrificed after 1 h, and the blood is collected by cardiac puncture. 7. Degerm mouse hair with 70% ethanol and open the thoracic cavity with sterile forceps and surgical scissors. 8. Insert the needle through the apex into the heart (see Note 20). 9. Apply gentle negative pressure to the syringe plunger. When blood appears in the syringe, the plunger should be gradually pulled back to obtain the maximum volume of blood. 10. The blood should be treated with 10 μl of EDTA (60 mg/ml) and centrifuged at 550  g for 10 min in a 10 mm radius rotor. 11. Approximately 500 μl of plasma should be harvested from each mouse. 12. The FI of the plasma FITC-dextran should be determined at an excitation wavelength of 485 nm and an emission wavelength of 528 nm using a plate reader.

4

Notes 1. Vessel permeability is highly dependent on the age and weight of the animal, so when comparing different mouse strains, the mice or other test subjects must have close to identical birth dates and weight [3]. 2. The endothelial barrier is also influenced by environmental conditions such as temperature, humidity, and the handling stress of the mouse; hence proper care should be taken to minimize these variations [3]. 3. Use 3% isoflurane for inducing anesthesia until the righting reflex is lost and the mouse is unresponsive to external stimuli, whereas 1.5% isoflurane can be used for anesthesia maintenance and ensure that the mouse has constant respiratory rates [16]. 4. If required, filter and sterilize the solution to remove any particulate matter that has not dissolved [3]. 5. Avoid all air bubbles that might have escaped into the syringe [3].

Vascular Permeability Assays In Vivo

173

6. Place the mouse restraint device on its side so the lateral tail vein is easily visible and is facing upward [3]. 7. The tail-vein injection technique should be mastered before starting the experiment [3]. 8. Before injection, apply pressure at the base of the tail so it works like a tourniquet [3]. 9. Do not apply back pressure to confirm proper placement as this might collapse the vein and observe the ease with which the plunger advances, as this is the proof of correct placement of the needle in the vein [3]. 10. A small drop of blood appears at the injection site if the needle is correctly placed [3]. 11. Perfuse each mouse with 10 ml saline via the left ventricle (after venting the left atrium [14]. 12. Isolate ear punches of 8 mm diameter using a biopsy punch and immediately keep on ice until mixed with formamide [14]. 13. Read absorbance at 610 nm for 50-μl triplicates in a 96-well plate [14]. 14. Due to the variety of factors that can impact the outcome of the experiment, it is always recommended to repeat the experiment for at least three times to perform the statistical analysis [3]. 15. Avertin solution (250 mg/kg body weight) can be injected by intraperitoneal route [21]. 16. LPS is an endotoxin, obtained from the outer membrane of gram-negative bacteria, used in studies to induced immune and inflammatory responses in mammals [21]. 17. LPS (0.5 mg/kg body weight) should be dissolved in 50 μl of PBS [21]. 18. 10 mg/kg body weight FITC-dextran should be added in 50 μl of sterile PBS [21]. 19. Hold mouse flat in hand and pipette FITC-dextran slowly but steadily onto each nostril. Pause pipetting if the mouse does not inhale the fluid and resume after complete fluid is inhaled [21]. 20. A 22-gauge needle attached to a 1-ml syringe should be used [21].

Acknowledgments Funds were provided by the NHLBI grant R01HL103952, NCATS grant UL1TR002378, Wilson Pharmacy Foundation (intramural), and Translational Research Initiative grant (intramural) to PRS.

174

Mir S. Adil and Payaningal R. Somanath

References 1. Bates DO (2010) Vascular endothelial growth factors and vascular permeability. Cardiovasc Res 87(2):262–271. https://doi.org/10. 1093/cvr/cvq105 2. Gao F, Artham S, Sabbineni H et al (2016) Akt1 promotes stimuli-induced endothelialbarrier protection through FoxO-mediated tight-junction protein turnover. Cell Mol Life Sci 73(20):3917–3933. https://doi.org/10. 1007/s00018-016-2232-z 3. Radu M, Chernoff J (2013) An in vivo assay to test blood vessel permeability. J Vis Exp 73: e50062. https://doi.org/10.3791/50062 4. Ashina K, Tsubosaka Y, Kobayashi K, Omori K, Murata T (2015) VEGF-induced blood flow increase causes vascular hyper-permeability in vivo. Biochem Biophys Res Commun 464 (2):590–595. https://doi.org/10.1016/j. bbrc.2015.07.014 5. McMurdy J, Reichner J, Mathews Z, Markey M, Intwala S, Crawford G (2009) Broadband reflectance spectroscopy for establishing a quantitative metric of vascular leak using the miles assay. J Biomed Opt 14 (5):054012. https://doi.org/10.1117/1. 3233654 6. Schniering J, Borgna F, Siwowska K et al (2018) In vivo labeling of plasma proteins for imaging of enhanced vascular permeability in the lungs. Mol Pharm 15(11):4995–5004. https://doi.org/10.1021/acs. molpharmaceut.8b00606 7. Chen J, Somanath PR, Razorenova O et al (2005) Akt1 regulates pathological angiogenesis, vascular maturation and permeability in vivo. Nat Med 11(11):1188–1196. https://doi.org/10.1038/nm1307 8. Bates DO, Hillman NJ, Williams B, Neal CR, Pocock TM (2002) Regulation of microvascular permeability by vascular endothelial growth factors. J Anat 200(6):581–597. https://doi. org/10.1046/j.1469-7580.2002.00066.x 9. Gao F, Sabbineni H, Artham S, Somanath PR (2017) Modulation of long-term endothelialbarrier integrity is conditional to the cross-talk between Akt and Src signaling. J Cell Physiol 232(10):2599–2609. https://doi.org/10. 1002/jcp.25791

10. Alwhaibi A, Verma A, Adil MS, Somanath PR (2019) The unconventional role of Akt1 in the advanced cancers and in diabetes-promoted carcinogenesis. Pharmacol Res 145:104270. https://doi.org/10.1016/j.phrs.2019. 104270 11. Gao F, Alwhaibi A, Artham S, Verma A, Somanath PR (2018) Endothelial Akt1 loss promotes prostate cancer metastasis via β-cateninregulated tight-junction protein turnover. Br J Cancer 118(11):1464–1475. https://doi.org/ 10.1038/s41416-018-0110-1 12. Artham S, Gao F, Verma A et al (2019) Endothelial stromelysin1 regulation by the forkhead box-O transcription factors is crucial in the exudative phase of acute lung injury. Pharmacol Res 141:249–263. https://doi.org/10. 1016/j.phrs.2019.01.006 13. Artham S, Verma A, Alwhaibi A et al (2020) Delayed Akt suppression in the lipopolysaccharide-induced acute lung injury promotes resolution that is associated with enhanced effector regulatory T cells. Am J Physiol Lung Cell Mol Physiol 318(4): L750–L761. https://doi.org/10.1152/ ajplung.00251.2019 14. Weis SM (2008) Chapter 5. Evaluating vascular leak in vivo. Methods Enzymol 444:99–114. https://doi.org/10.1016/S0076-6879(08) 02805-X 15. Islam MT, Tasciotti E, Righetti R (2020) Estimation of vascular permeability in irregularly shaped cancers using ultrasound Poroelastography. IEEE Trans Biomed Eng 67 (4):1083–1096. https://doi.org/10.1109/ TBME.2019.2929134 16. Brash JT, Ruhrberg C, Fantin A (2018) Evaluating vascular Hyperpermeability-inducing agents in the skin with the miles assay. J Vis Exp 136:57524. https://doi.org/10.3791/ 57524 17. McClure N, Robertson DM, Heyward P, Healy DL (1994) Image analysis quantification of the miles assay. J Pharmacol Toxicol Methods 32 (1):49–52. https://doi.org/10.1016/10568719(94)90017-5 18. Ferrero ME (2004) In vivo vascular leakage assay. Methods Mol Med 98:191–198.

Vascular Permeability Assays In Vivo https://doi.org/10.1385/1-59259-7718:191 19. Natarajan R, Northrop N, Yamamoto B (2017) Fluorescein isothiocyanate (FITC)-dextran extravasation as a measure of blood-brain barrier permeability. Curr Protoc Neurosci 79:9581–9585. https://doi.org/10.1002/ cpns.25 20. Boutoille D, Marechal X, Pichenot M, Chemani C, Guery B, Faure K (2009) FITCalbumin as a marker for assessment of

175

endothelial permeability in mice: comparison with 125I-albumin. Exp Lung Res 35 (4):263–271. https://doi.org/10.1080/ 01902140802632290 21. Chen H, Wu S, Lu R, Zhang YG, Zheng Y, Sun J (2014) Pulmonary permeability assessed by fluorescent-labeled dextran instilled intranasally into mice with LPS-induced acute lung injury. PLoS One 9(7):e101925. https://doi. org/10.1371/journal.pone.0101925

Methods in Molecular Biology (2021) 2367: 177–191 DOI 10.1007/7651_2020_309 © Springer Science+Business Media New York 2020 Published online: 21 July 2020

Endothelial Permeability Assays In Vitro Mir S. Adil and Payaningal R. Somanath Abstract The endothelium is a thin layer of squamous cells that acts as a semipermeable barrier regulating vascular permeability to let molecules pass through it thereby maintaining tissue fluid homeostasis. Physiological increase in endothelial or vascular permeability is transient, transpired by post-tissue injury during the initial phases of healing, whereas pathological permeability is persistent commonly witnessed in conditions such as atherosclerosis, chronic inflammation, tumor growth, and diabetic retinopathy. The in vivo or in situ use of animal models in the assessment of permeability not only raises inevitable ethical concerns but also confers difficulty to apply to high-throughput screening. Therefore, there is an ever-increasing dependency on in vitro studies to assess drug permeability, and various research programs have suffered to develop appropriate in vitro assays for measurement and prediction. In vitro models that both mimic in vivo microvascular endothelium and can be utilized to record changes in endothelial permeability are vital in delineating the mechanisms involved in the prevention and treatment of disorders related to vascular permeability. The Transwell® and the electric cell-substrate impedance sensing (ECIS) assays are extensively used to assess the trans-endothelial permeability of solutes such as albumin, dextrans, and sucrose across endothelial monolayers and based on electrical resistance, etc. These models have several advantages such as the ease to perform and avoid the complexities of using a live animal. Keywords Cell barrier, In vitro permeability, Endothelial cells, Transwell® assay

1

Introduction A thin layer of vascular endothelium lines the inner part of blood vessels to maintain vascular integrity [1, 2]. It acts as a semipermeable barrier regulating selective vascular permeability to let molecules pass through it thereby maintaining tissue fluid homeostasis [1, 3–5].

1.1 Vascular Permeability

Important differences lie between the basal vascular permeability and the blood-tissue barrier modulations that occur during pathological states. These include acute or chronic phases of hyperpermeability, the barrier-crossing anatomic pathways followed by solutes, and the composition of extravasate [5]. The physiological increase in endothelial permeability is transient that is transpired post-tissue injury during the initial phases of healing. In contrast, pathological endothelial permeability is persistent and has been implicated in disease states such as

177

178

Mir S. Adil and Payaningal R. Somanath

atherosclerosis, chronic inflammation, tumor growth, diabetic retinopathy, etc. [6]. Failure of vascular barrier integrity has been reported to result in serious outcomes including hemorrhage, edema, inflammation, and tissue ischemia [2]. 1.2 Physiological State of EndothelialBarrier

Endothelial cells are interlinked by a complex set of junction proteins that include adherens junctions (AJs) and tight junctions (TJs) [2, 3]. While vascular endothelial cadherin (VE-cadherin) is the major structural protein in the AJs [7], occludins and claudins are the TJ proteins linked to the protein zona occludens (ZO-1/2/3) critical in maintaining the permeability and integrity [2]. Under normal conditions, inter-endothelial junctions open to permit the passage of molecules and surveillance cells in a dynamic and sizeselective manner [3].

1.3 Pathophysiological State of EndothelialBarrier

Under pathophysiologic conditions, the integrity of the vascular endothelium can be disturbed, thereby leading to fluid leakage into the extravascular space. Damage to TJs is thought to be an integral component of pathophysiologies related to vessel injury [2]. When it comes to the tumor microenvironment, vascular endothelium of the tumor cells not only plays a critical role in tumor angiogenesis, perfusion, and metastasis but also acts as the foremost defense in a patient’s fight against the tumor metastasis to other significant organs of the body [8].

1.4 Routes of Transportation

Plasma proteins and solutes transportation or leukocyte transmigration through the semipermeable vascular endothelium layer can take place by two different routes, namely, transcellular and paracellular. While the former involves caveolae-mediated vesicular transport, the latter makes use of the space between interendothelial junctions [4]. Caveolae, vesiculo-vacuolar organelles (VVOs), and fenestrae in the transcellular pathway allow molecules to cross the endothelial cells. The size of vesicles or vacuoles of VVOs ranges from caveolaesized vesicles to vacuoles with as much as tenfold larger volumes. The homeostatic transport of proteins and lipids occurs largely via transcellular routes, whereas pathological vascular leakage usually takes place by paracellular routes. Cells and macromolecules with size larger than 3 nm escape the blood vessel through the paracellular pathway, which is facilitated by the coordinated opening and closing of the cell-cell junctions in endothelium [5].

1.5 Regulators of Vascular Permeability

VEGF is a vital regulator of vascular permeability as its activation results in phosphorylation and disassembly of VE-cadherin thereby compromising the barrier integrity [5]. Other inflammatory factors such as histamine or thrombin also induce a transient and non-pathological increase of vascular permeability [1]. Pro-inflammatory cytokines such as tumor necrosis factor

Endothelial Permeability Assays In Vitro

179

(TNF-α) and interleukin-1β (IL-1β) have shown to increase the endothelial permeability by disrupting the expression of VE-cadherin and ZO-1 [2]. In contrast to the aforesaid agents, angiopoietin-1 enhances endothelial-barrier function, and its blocking leads to disruption of the endothelial barrier [5, 9]. Ras-related GTP-binding proteins show varied effects as Rho A can lead to the disruption of barrier function through Rac1 activation, whereas Cdc42, when activated, stabilizes cell-cell junction. The TGFβ also shows mixed effects as its signaling through ALK5 leads to stabilized vascular endothelium, whereas ALK1-mediated pathway limits ALK5 signaling, leading to vascular destabilization [5]. Akt1, an isoform of a serine-threonine kinase Akt [10], was found to be essential for vascular endothelial-barrier integrity [11]. Endothelial-specific knockdown of Akt1 led to increased vascular permeability via FoxO- and β-catenin-mediated repression of endothelial tight-junction claudin expression, mainly claudin-5 [8]. Moreover, the depletion of Akt1 has shown to prevent the restoration of basal endothelial-barrier resistance post-VEGFinduced permeability and impair angiopoietin-1-mediated endothelial-barrier enhancement in vitro and vascular permeability in vivo [9]. 1.6 Significance of Vascular Permeability Studies

Permeability is a vital factor contributing to the pharmacokinetic aspects of compounds that include absorption, distribution, metabolism, and excretion [12]. The in vivo or in situ use of animal models in the assessment of permeability not only raises inevitable ethical concerns but also confers difficulty to apply to highthroughput screening. Therefore, there is an ever-increasing dependency on in vitro studies to assess drug permeability, and various research programs have suffered to develop appropriate in vitro assays for measurement and prediction [12]. Given the complex manipulations requirement to study molecular or pharmacological barrier function in vivo, cultured endothelial cells are frequently used to replicate the microvascular endothelium, and most of these involve confluent endothelial cells cultured on a porous membrane filter in Transwell® chambers [5]. In vitro models that both mimic in vivo microvascular endothelium and can be utilized to record changes in endothelial permeability are vital in delineating the mechanisms involved in the prevention and treatment of disorders related to vascular permeability [6]. The methods to study vascular permeability have been of interest for many years as it is an essential characteristic of the vascular system [5, 13]. The very first set of methods developed to assess and record vascular permeability mainly relied on in vivo assays, such as the Miles assay [14] or ex vivo assays that involve the use of blood vessels isolated from the organism [10]. Because of the

180

Mir S. Adil and Payaningal R. Somanath

challenges in handling these models, the in vitro model, Transwell® assay, was developed which allows determining the permeability of a monolayer of endothelial cells with ease [1]. 1.7 In Vitro Permeability Assay in Transwell® Plates

Measuring barrier characteristics of interconnected cell layers and their dynamic variations in vitro looks back on a long history [15]. Over the years, the Transwell® assay is extensively used to assess the trans-endothelial permeability of solutes such as albumin, dextrans, and sucrose across endothelial monolayers. These models have several advantages such as the ease to perform and avoid the complexities of using a live animal [5].

1.8 Limitations of Transwell® In Vitro Permeability Assay

In vitro permeability assays do not come without drawbacks. Some of the limitations are as follows: 1. The monolayers formed onto polycarbonate membranes or on matrix-coated membranes in the in vitro assays are not as tight as their in vivo counterparts primarily due to the lack of basement membrane [5, 6] and absence of other types of cells such as pericytes or smooth muscle cells. Also, structures that are vital for macromolecular extravasation are not present at the in vitro level [5]. 2. The model lacks the three-dimensional (3D) structure of a blood vessel. Although recently in vitro models based on electrical resistance such as electrical cell-substrate impedance sensing (ECIS) were developed to overcome this drawback, they do not let the investigator know if a protein crossed the barrier [1]. 3. The source of endothelial cells in the in vitro models is primary cells that originated from umbilical veins or cell lines with different characteristics than the microvessels [6].

1.9 Electric CellSubstrate Impedance Sensing (ECIS) Assay to Measure EC-Barrier Function

ECIS biosensor technology, developed by Applied BioPhysics (USA), has a broad range of applications based on the cell type of interest [17]. It is a piece of versatile equipment that is capable of measuring as well as monitoring the structural and functional properties of live cells in a real-time fashion [17–19]. It relies on the alterations in the impedance of electrode-cell interface due to adhesion and motility of anchorage-dependent cells [20]. Traditionally, intensive labor is involved in cell attachment and spreading measurements along with a lot of manipulations of the cell cultures for microscopic evaluation of its behavior. ECIS can measure these in a much easier and more convenient way [19]. The process involves the inoculation of living cells on an array consisting of eight separate culture wells [17]. The cells are then allowed to attach and grow on the small gold electrode deposited on the bottom of the well [19]. A weak alternating current (10–105 Hz) is applied through the electrode array to provide a means of measuring the ability of cells grown in a monolayer to

Endothelial Permeability Assays In Vitro

181

Fig. 1 Endothelial cell monolayer without any space in between prevents the passage of electric current, and high resistance is recorded (above). Space between cells allows the passage of electric current, and low resistance is recorded (below)

impede the movement of electrons through and between individual cells. The impedance measurement provides information on two markers of cell behavior, resistance (R) and capacitance (C). While resistance is indicative of the barrier function of the cells, capacitance provides measurements related to the overall coverage of the electrode by the cell monolayer [17]. The multifrequency provision in ECIS allows the impedance data to be mathematically modeled thereby indicating the contribution of the paracellular junctional space, the basal adhesion of the cells, and the cell membrane to the total impedance to current. Low frequency restricts the movement of current between the cells in presence of intercellular junctions (as depicted in Fig. 1), whereas high frequencies (>10,000 Hz) allow the current to flow through the cell body owing to low reactive capacitance, where the capacitive function of the cell membrane is indicative of the extent of cell coverage over the electrode [17].

2

Materials

2.1 In Vitro Permeability Assay 2.1.1 Cell Culture [6, 13, 16]

Primary human endothelial cells (ECs) or the desired cell line is required along with the following: 1. Appropriate growth medium: e.g., EBM-2 medium with a GM-2 Bullet Kit from Lonza for Human Lung ECs (HLECs) 2. Cell detachment buffer (e.g., 0.05% trypsin) 3. Sterile phosphate-buffered saline (PBS) 4. Sterile cell culture hood 5. Pipettors, liquid aspirators, etc. for the handling of cells and liquid reagents

182

Mir S. Adil and Payaningal R. Somanath

6. Sterile cell culture flasks, centrifuge tubes, pipettes, its tips, etc. for the handling of cells 7. Sterile CO2 tissue culture incubator 8. 70% ethanol to disinfect forceps, surfaces, plasticware, etc. 9. Low-speed centrifuge for cell harvesting 10. Hemocytometer for cell count 11. Trypan blue or equivalent viability stain 12. Microscope 2.1.2 Transwell® Assay [6, 13, 16]

In in vitro vascular permeability, the assay is performed in a 24-well receiver plate with 24 individual hanging cell culture inserts. The inserts contain minute pores within a transparent polyethylene terephthalate (PET) membrane as shown in Fig. 2a. The following materials are required to proceed with the assay: 1. Cell culture permeable inserts, 24 well 2. Cell culture permeable insert receiver plates, 24 well 3. Matrigel basement membrane matrix 4. Tris, sodium chloride (coating buffer) 5. 0.2 μm filter unit 6. Sterile forceps 7. Positive displacement pipette 8. Syringes 9. Fluorescein isothiocyanate (FITC) – dextran 10. Plate shaker 11. Black opaque plate, 96 well 12. A fluorescence plate reader with filters for 485 nm and 535 nm excitation and emission, respectively (appropriate for FITC/ fluorescein signal) 13. 0.5% crystal violet stain

2.2 ECIS Assay [21, 22]

1. An ECIS instrument placed in a sterile CO2 tissue culture incubator and connected to a computer installed with a compatible software. 2. ECIS culture well array: choose an array based on the application as various types of arrays are available that helps in determining wound healing migration, micromotion assays, or barrier function/permeability assays. Use 8W10E+ array for assessing the endothelial barrier integrity. 3. 10 mM sterile solution of L-cysteine in distilled water (commercially available as ECIS electrode-stabilizing solution) 4. Petri dish.

Endothelial Permeability Assays In Vitro

183

Fig. 2 Diagrammatic representation of the Transwell® setup and the steps involved in the seeding of cells and assay

5. Microscope. 6. Materials required for preparation of cell suspension (as discussed above).

3

Methods

3.1 In Vitro Permeability Assay 3.1.1 Endothelial Cell Culture [13]

1. Activate ECs or desired cell lines under a sterile cell culture hood and grow in an appropriate medium (EBM-2 with a GM-2 Bullet Kit for HLECs). 2. Place them in a humidified 5% CO2 incubator at 37
C and routinely passage when the confluency of 80–90% is achieved (see Note 1). 3. Visually inspect cells and wash them with 4 ml sterile PBS (for 75 cm2 flask) before harvest.

184

Mir S. Adil and Payaningal R. Somanath

4. Add 4 ml 0.05% trypsin and incubate at room temperature for 1–3 min or until cells start to detach. 5. Add 4–6 ml growth medium or trypsin neutralizing solution to inactivate the cell detachment buffer. 6. Gently pipette the cells out of the flask into a sterile centrifuge tube. 7. Gently mix the cell suspension, remove a sample, and dilute it with trypan blue in a 1:1 ratio before counting on a hemocytometer to check its viability. 8. Centrifuge cells to pellet (1000  rpm, 3–4 min). 9. Gently resuspend the pellet in growth medium to a concentration of 0.5–2  106 cells/ml. 10. Additional compounds such as cytokines, pharmacological agents, etc. can be added to the cell suspension if pretreatment of the endothelial cells is required. 3.1.2 Coating the Transwell® Plates [6, 13]

The following steps must be performed using the aseptic technique: 1. Matrigel matrix should be reconstituted, aliquoted, and stored as required. 2. Matrigel matrix along with the coating buffer must be kept ice-cold during the entire procedure. 3. Prepare a coating buffer by mixing 0.01 M Tris (pH 8.0) with 0.7% NaCl followed by filtration using a 0.2 μm sterile filter unit. 4. Thaw Matrigel matrix aliquot on the ice at 4
C and swirl the vial to ensure the content inside is evenly dispersed. 5. Keep the product on the ice and handle using sterile technique. 6. Allow the coating buffer to cool for 2 h in an ice bath in a cold room or refrigerator. 7. Pipettes, syringes, or containers intended to be used for handling the Matrigel matrix must be chilled before use. 8. Syringes and pipettes can safely be placed on ice if placed within a plastic bag. 9. Prepare a coating solution by mixing the Matrigel matrix (final concentration of 200–300 μg/ml) with coating buffer in a final volume of 2 ml which would be enough to coat 24 inserts. 10. Thoroughly mix the coating solution containing the Matrigel matrix by gently swirling followed by placing the tube on ice. 11. For each new syringe, fill halfway with Matrigel matrix, expel, and then fill to coat.

Endothelial Permeability Assays In Vitro

185

12. If using a positive displacement pipet, a larger volume of Matrigel matrix will be needed to prevent or minimize bubble formation. 13. Based on the treatment in the study, select the Transwell® insert pore size which ranges from 0.4 to 8.0 μm. 14. Under the hood, remove the lid from the Transwell® plate receiver with inserts. 15. Use a sterile syringe or pipette to carefully add 0.1 ml of the diluted Matrigel matrix coating solution to each permeable insert. 16. Minimize contact of the Matrigel matrix with the sidewalls of the permeable support. 17. Repeat the coating process with the remaining permeable supports. 18. Incubate plates with the coated permeable inserts (upper chambers) at 37
C for 2 h. 19. Carefully remove the remaining liquid from the insert (see Note 2). 20. The coated invasion chambers are now ready for use. 3.1.3 Seeding ECs into Transwell® Plates [6, 16]

1. Deposit 100 μl of the diluted cell suspension with ~105 cells (see Note 3) into the Transwell® insert on top of the Matrigelcoated filter as represented in Fig. 2b (see Note 4). 2. Let the cells adhere by incubating the plate for 30–60 min at 37
C (Fig. 2c). 3. Add 200 μl growth medium to the insert. 4. Hold insert with forceps (see Note 5) and add 1 ml medium to the receiver well (in the 24-well plate), as diagrammatically represented in Fig. 2d (see Note 6). 5. Incubate at 37
C for 24–48 h (see Note 7). 6. Cells can be treated and incubated further as per the requirement which could lead to strengthening or disruption of the barrier as represented in Fig. 2e.

3.1.4 Transwell® Assay Protocol [6, 16]

1. Following the completion of treatment, transfer each insert into a new receiver well on a fresh plate (Fig. 2f). 2. Add 1 ml of growth medium to each receiver well wherever the insert is being placed to be analyzed. 3. Remove the medium from the insert without disturbing the cell monolayer (see Notes 2 and 8). 4. Prepare a working solution of FITC-dextran in medium to make a final dilution of 10 μg/ml (see Note 9). 5. Make a sufficient volume of solution to add 150 μl per insert.

186

Mir S. Adil and Payaningal R. Somanath

6. Add 150 μl of FITC-dextran working solution to each insert as denoted in Fig. 2g. 7. Incubate the plate, protected from light, for 20 min at room temperature. (This permeation time is suggested, although the duration can be optimized by the end user). 8. During this time, the FITC-dextran molecules pass through the endothelial monolayer depending on the integrity of the barrier (Fig. 2h). 9. To avoid any inhomogeneous FITC-dextran distribution in the lower compartment, Transwell® plates can be gently and repetitively shaken. 10. Stop permeation by removing the inserts from the wells as shown in Fig. 2i. 11. Thoroughly mix the medium in the wells of the receiver tray (now containing FITC-dextran that crossed the monolayer). 12. Inserts may be temporarily held in a previous receiver tray that has been emptied of liquid. 13. Remove 100 μl of the medium from each well of the receiver well and transfer to wells of a black 96-well opaque plate to perform fluorescence measurement. 14. The removed volume should be replaced by a fresh medium. 15. Medium from the receiver well can be collected at multiple time points and eventually transferred to the opaque plate. 16. Read the plate using a fluorescence plate reader with filters appropriate for 485 nm and 535 nm excitation and emission, respectively (or similar FITC/fluorescein-compatible wavelengths) (see Note 10). 17. The mean fluorescence recorded from the receiver well of untreated cells at the final time point (30 or 60 min) can be set as 1. Data are expressed as relative changes compared to control levels (see Note 11). 18. The absolute permeability P [cm/s] can be calculated by the following equation: P ¼ ½CðtÞ  Cðt0 Þ  V=A  t  C0 where, l

l

l

C(t) is the concentration [μg/ml] of FITC-dextran in the samples that were taken from the receiver wells after 30, 60, or 90 min. C(t0) is the FITC-dextran concentration [μg/ml] of the samples taken after 0 min. t is the duration of the flux (s).

Endothelial Permeability Assays In Vitro l

l

187

V is the volume [cm3] in the lower compartment; A is the surface of the Transwell® membrane [cm2]. C0 is the initial concentration [μg/ml] of the tracer on the donor side.

19. The concentration of FITC-dextran in each sample is determined by reference to a FITC-dextran standard curve. 3.1.5 Monolayer Staining [16]

1. After completion of FITC-dextran permeability testing, the endothelial monolayer can be stained to obtain brightfield images to study monolayer integrity. 2. This can also be performed on a spare cell-seeded insert following the 72 h monolayer formation time to check for monolayer confluency before starvation or permeability treatment. 3. However, stained inserts cannot be used for FITC-dextran permeability testing or further cell culture; sample numbers should be planned to account for discarding of inserts stained before permeability testing. 4. Place the insert in an empty receiver plate well with no liquid; remove the medium carefully without disturbing the cell monolayer (see Notes 2 and 8). 5. Add 100 μl of 0.5% crystal violet staining solution to the insert, cover the plate, and incubate at room temperature for 20 min. 6. Carefully remove the cell stain and discard it as hazardous waste. 7. Rinse the insert twice with PBS, filling the insert and receiver plate well with 200 μl and 1 ml of buffer, respectively, for each rinse. The insert may be left in the second rinse during microscopic (brightfield) imaging.

3.2 ECIS Protocol for EC-Barrier Impedance Assay [21, 22]

1. Flood each well with ~200 μl of L-cysteine solution and aspirate the solution after keeping the array for 10 min at room temperature. 2. Rinse at least twice with sterile distilled water. 3. The wells can be coated with proteins (see Note 12) and rinsed well (if desired), or 200 μl medium can be added directly to each well for immediate use. 4. Place the array in a Petri dish (to lower the risk of contamination) and transfer it to the incubator. 5. Remove the array from the Petri dish and connect it to the holder of the station. 6. Click “Setup/Check” in the software to ensure the array is properly connected.

188

Mir S. Adil and Payaningal R. Somanath

7. While green light indicates a properly connected array, red light signifies any interruption between the array and the station (see Note 13). 8. Remove the array from the station and place it in the Petri dish before transferring it to the hood. 9. Rinse out the medium from the wells and add 400 μl of cell suspension with optimal cell count (see Note 14). 10. Incubate the array for 24 h or until the desired confluency is achieved (see Note 15). 11. Aspirate the medium by touching the wall of the array well and add 200 μl of the fresh medium before connecting the array to the station. 12. Click the “Start” button on the software and name the file to begin the experiment (choose the desired frequency for the experiment, or “multiple frequencies” can also be selected). 13. Let the cells adjust and equilibrate to the experimental conditions before adding any treatment. 14. Prepare the compound to be tested at 2 the final desired concentration in the same solution (medium) as is in the wells and equilibrated to the incubator temperature (see Notes 16 and 17). 15. After 1–2 h, “Pause” the experiment and add 200 μl of prepared drug or inhibitor-containing medium to the respective wells without disturbing the cell layer and “Resume” the experiment (see Notes 18 and 19). 16. Click the “Finish” button once the desired data is collected. 17. Press “Pause” if the array plate must be removed from the holder during the experiment (see Note 20). 18. Click the “Analyze Data” button and save it in the form of graphical image and/or export data in the Microsoft Excel file. 19. Multiple wells can be grouped based on the treatment to obtain the average values (see Note 21). 20. Data can also be normalized if the researcher wants to use the zero time as the reference time.

4

Notes 1. Use cells between passages four and nine as more passaging make cells form a leaky monolayer without any treatment [6]. 2. Be careful not to touch the insert membrane at times as this may damage the monolayer integrity of the underlying porous membrane. During removal or addition of liquid, touch the insert wall with the pipette without disturbing the base [16].

Endothelial Permeability Assays In Vitro

189

3. To determine optimal seeding density on a porous membrane, seed cells in a range that brackets the typical density used for nonporous membrane culture [13]. 4. Transwell® inserts with a wrinkled appearance should not be used as these are prone to leak [6]. 5. Sterilize forceps with 70% ethanol before handling inserts [16]. 6. Ensure medium completely covers the underside of the insert and that no bubbles are present as air may get trapped at the interface [16]. 7. Let the cells incubate for a longer period and/or use freshly prepared medium if you notice sub-confluent endothelial monolayer. Endothelial survival factors such as Ang-1 and VEGF can be used to improve the confluency of monolayer [16]. 8. Empty and refill one insert at a time. Make sure some residual liquid is always present on top of the insert membrane and try to minimize the time required for aspirating or filling the insert to avoid drying of the cell monolayer [16]. 9. Always protect the FITC-dextran solution from light [16]. 10. Adjust the concentration and/or permeation time of FITCdextran if you experience a very high/very low fluorescent signal [16]. 11. In the case of no/minimal effects on treatment, the composition of the medium can be modulated to detect effects resulting from various factors [16]. 12. If pre-coating is desired, avoid phosphate buffer to prepare protein as it can interfere with the adsorption of some proteins [21]. 13. If you see a red-light indicator for any well after array insertion, adjust the placement of the array and click “Setup” again [21]. 14. Try to avoid clumping of cells and ensure accurate cell count before adding the cell suspension as unequal cell distribution might lead to variations in results [22]. 15. Make sure the temperature of the array and cell suspension added to the wells is close to that of the incubator as colder cell suspension leads to heating of array wells from the bottom eventually reducing the density of cells in the central region [21]. 16. Calculate the drug or inhibitors required for 400 μl of medium per well and add the required volume to 200 μl of medium per well as the remaining 200 μl of medium is already present in the array wells attached to the station [21]. 17. In addition to treatments, be certain to include a control without the compound [21].

190

Mir S. Adil and Payaningal R. Somanath

18. Even if the compound addition is carried out without removing the array, press the “Pause” button to avoid any unwanted impedance spike [22]. 19. Do not exceed 400 μl of medium per well at any time (for eight-well array plate) [21]. 20. Try not to remove the plate from the holder during the experiment. If the need arises, work quickly so temperature drops are minimized [22]. 21. For a lucid representation, the colors of the eight-well arrays can be customized by selecting a different colormap from the menu [22].

Acknowledgments Funds were provided by the NHLBI grant R01HL103952, NCATS grant UL1TR002378, Wilson Pharmacy Foundation (intramural), and Translational Research Initiative grant (intramural) to PRS. References 1. Pauty J, Usuba R, Takahashi H et al (2017) A vascular permeability assay using an in vitro human microvessel model mimicking the inflammatory condition. Nano 1(1):103–113 2. Tang X, Di X, Liu Y (2017) Protective effects of donepezil against endothelial permeability. Eur J Pharmacol 811:60–65 3. Benelhaj NE, Maraveyas A, Featherby S, Collier MEW, Johnson MJ, Ettelaie C (2019) Alteration in endothelial permeability occurs in response to the activation of PAR2 by factor Xa but not directly by the TF-factor VIIa complex. Thromb Res 175:13–20 4. Di A, Mehta D, Malik AB (2016) ROS-activated calcium signaling mechanisms regulating endothelial barrier function. Cell Calcium 60(3):163–171 5. Park-Windhol C, D’Amore PA (2016) Disorders of vascular permeability. Annu Rev Pathol 11:251–281 6. Martins-Green M, Petreaca M, Yao M (2008) An assay system for in vitro detection of permeability in human "endothelium". Methods Enzymol 443:137–153 7. Navarro P, Caveda L, Breviario F, Maˆndoteanu I, Lampugnani MG, Dejana E (1995) Catenin-dependent and -independent functions of vascular endothelial cadherin. J Biol Chem 270(52):30965–30972

8. Gao F, Alwhaibi A, Artham S, Verma A, Somanath PR (2018) Endothelial Akt1 loss promotes prostate cancer metastasis via β-cateninregulated tight-junction protein turnover. Br J Cancer 118(11):1464–1475. https://doi.org/ 10.1038/s41416-018-0110-1 9. Gao F, Artham S, Sabbineni H et al (2016) Akt1 promotes stimuli-induced endothelialbarrier protection through FoxO-mediated tight-junction protein turnover. Cell Mol Life Sci 73(20):3917–3933 10. Alwhaibi A, Verma A, Adil MS, Somanath PR (2019) The unconventional role of Akt1 in the advanced cancers and in diabetes-promoted carcinogenesis. Pharmacol Res 145:104270 11. Artham S, Verma A, Alwhaibi A et al (2020) Delayed Akt suppression in the lipopolysaccharide-induced acute lung injury promotes resolution that is associated with enhanced effector regulatory T-cells. Am J Physiol Lung Cell Mol Physiol 318(4): L750–L761 12. Lee JB, Son SH, Park MC et al (2015) A novel in vitro permeability assay using a threedimensional cell culture system. J Biotechnol 205:93–100 13. Islam S, Flaherty P (2018) Assay methods protocol: cell invasion assay. Corning Incorporated, Life Sciences Tewksbury, MA USA

Endothelial Permeability Assays In Vitro 14. Bischoff I, Hornburger MC, Mayer BA, Beyerle A, Wegener J, Fu¨rst R (2016) Pitfalls in assessing microvascular endothelial barrier function: impedance-based devices versus the classic macromolecular tracer assay. Sci Rep 6:23671 15. Wegener J (2019) Measuring the permeability of endothelial cell monolayers: teaching new tricks to an old dog. Biophys J 116 (8):1377–1379 16. Millipore (2010) In vitro vascular permeability assay (24-well) catalog no. ECM644. Revision A 17. Robilliard LD, Kho DT, Johnson RH, Anchan A, O’Carroll SJ, Graham ES (2018) The importance of multifrequency impedance sensing of endothelial barrier formation using ECIS Technology for the Generation of a strong and durable Paracellular barrier. Biosensors (Basel) 8(3):64

191

18. Chiu SP, Lee YW, Wu LY et al (2019) Application of ECIS to assess FCCP-induced changes of MSC micromotion and wound healing migration. Sensors (Basel) 19(14):3210 19. Yang JM, Chen SW, Yang JH, Hsu CC, Wang JS (2016) A quantitative cell modeling and wound-healing analysis based on the electric cell-substrate impedance sensing (ECIS) method. Comput Biol Med 69:134–143 20. Rahman AR, Lo CM, Bhansali S (2009) A detailed model for high-frequency impedance characterization of ovarian cancer epithelial cell layer using ECIS electrodes. IEEE Trans Biomed Eng 56(2):485–492 21. ECIS User Handbook (2018) Applied BioPhysics, Inc., vol 18, pp 1–16 22. Electric cell-substrate impedance sensing. Applied BioPhysics Operation Manual for all ECIS Systems Version 1.2.123:1–86

Methods in Molecular Biology (2021) 2367: 193–205 DOI 10.1007/7651_2020_312 © Springer Science+Business Media New York 2020 Published online: 20 August 2020

Mapping Receptor Antibody Endocytosis and Trafficking in Brain Endothelial Cells Mikkel R. Holst, Simone S. E. Nielsen, and Morten S. Nielsen Abstract Drug delivery to the brain is a tremendous problem for the academic society and the industry. One solution with a huge potential is to use endocytic receptors as carriers. Here we describe how endocytic activity and subcellular trafficking of a specific receptor in brain endothelial cells can be characterized in three steps. (1) Labeling, endocytosis, and trafficking of a specific receptor at given time points in a pulse-chase experiment. (2) Fixed antibody labeling and co-staining of subcellular markers for image acquisition. (3) Analysis and quantification of co-localization between the receptor and subcellular markers in ImageJ. Keywords Brain endothelial cells, Receptor antibody trafficking, Endocytosis, Drug delivery, ImageJ, Co-localization, Immunofluorescence staining

1

Introduction Endocytosis is the cellular process by which extracellular components such as solutes, macromolecules, membrane receptors, etc. are internalized into cells. Micropinocytosis, phagocytosis, and pinocytosis are the three major types of classified endocytic mechanisms. Pinocytosis can be further subdivided into several different highly dynamic endocytic mechanisms such as clathrinmediated, caveolae-dependent, Arf6-dependent endocytosis, etc. [1]. It is via pinocytosis that membrane-bound receptors and its cargo are internalized. After internalization, receptors traffic to the early endosomes. Here, receptor interaction with different cytosolic proteins and membrane lipids will direct its trafficking to recycling endosomes, late endosomes, lysosomes, or trans-Golgi network (TGN) [2]. This endosomal trafficking system is often referred to as the endo-lysosomal system where coat-associated proteins participate in different phases of its maturation for regulation of receptor trafficking and cargo delivery. Antibody labeling of specific coatassociated proteins at various steps of the processing trafficking system can consequently be used to map the trafficking route of receptors in this system. Trafficking of receptors and its cargo in the endo-lysosomal system is fast, and the time a receptor uses to traffic

193

194

Mikkel R. Holst et al.

from the plasma membrane to, e.g., lysosomes, TGN or back to the cell surface can occur within few minutes. The endocytosis and subcellular receptor trafficking steps can be followed by live cell imaging, but the continuously uptake and labeling can make specific localization and quantification difficult. Consequently, a pulsechase technique providing fixed samples followed by immunofluorescence (IF) staining using subcellular markers providing snapshots of the localization is of high value to map receptormediated endocytosis and trafficking in brain endothelial cells (BECs). Saturation of a specific receptor at the cell membrane with a targeting antibody (the pulse step) can be done by lowering the temperature below 10
C whereby endocytic processes are blocked [3]. Heating the cells back to 37
C in a CO2 incubator will reinitiate the endocytic process whereby receptor trafficking can be chased in given time intervals in the endo-lysosomal system. Subsequent to this pulse-chase technique, IF staining, often referred to as immunocytochemistry (ICC), is used for visualizing specific target components in the cells. By specifying primary antibody affinities to antigens of interest and detecting these by conjugated fluorophores and fluorescent imaging, the method in general allows visualization of a broad array of targets with numerous applications within biomedical research, e.g., characteristics of cell surface and intracellular protein movements as in this assay. The primary antibodies are raised in a range of different species, e.g., mouse, rabbit, donkey, and goat, and can provide precise antigen specificities, with a large range of different protein targets being commercially available [4]. The IF technique includes both a direct and an indirect method. In direct IF, the fluorophore is conjugated directly to the primary antibody and represents the simplest IF procedure. This approach is suitable for studying receptor trafficking in living cells. In indirect IF, applied in this protocol, the primary antibodies are detected by use of fluorophore-conjugated secondary antibodies raised against the species of the primary antibody. Introducing a secondary antibody adds flexibility in the choice of fluorophore combinations to stain for multiple targets and results in an amplification of the fluorescent signal. However, there is a risk of species cross-reactivity, as the secondary antibodies may cross-react with species additional to the target species. However, such nonspecific binding is decreased by usage of pre-adsorbed secondary antibodies, which have been passed through a column matrix of immobilized serum proteins from putative cross-reactive species. Furthermore, biological samples may contain a high number of endogenous immunoglobulins, which the secondary antibody can bind, resulting in background signal. In order to evaluate nonspecific binding of the antibodies (background), an isotype control lacking specificity to the target should be included in a parallel (negative control) experiment. Although more circumstantial, the indirect IF method is the most prevalent method due to the high sensitivity and flexibility in the antibody combinations.

Mapping Receptor Antibody Endocytosis and Trafficking in Brain Endothelial Cells

195

There are a number of essential steps preliminary to the staining procedure with antibodies. The first step comprises cellular fixation, stopping cellular trafficking, immobilizing, and preserving the cellular morphology, which is crucial for the final staining result. Depending on the specific choice of primary antibodies in the subsequent staining procedure, different fixatives and processing methods are recommended. The most commonly used fixatives include the cross-linking reagent paraformaldehyde and the denaturing fixative methanol [5]. Depending on the used fixative, subsequent permeabilization of the lipid cellular membrane is necessary to allow specific antibodies access to intracellular antigens. This can be obtained by the use of lipid-soluble agents such as Triton X-100 or saponin. Furthermore, nonspecific binding sites must be blocked in order to prevent the antibodies binding sites not related to the specific antibody-antigen reactivity, e.g., due to charge or hydrophobicity reactions. Commonly used blocking agents include bovine serum albumin (BSA) or serum antibodies from the same species as the applied secondary antibody. Specific proteins of interest for the endo-lysosomal system may include markers such as EEA1 and Rab5 for early endosome, Rab7 for late endosome, Rab4 for recycling endosome, MPR300 for TGN, and Lamp1 for lysosomes [2]. Following incubation with primary and eventual secondary antibodies according to the manufacturer’s recommendations, these are visualized using antibodyconjugated fluorophores. Fluorophores, in the literature also referred to as fluorescent or reactive dyes, are fluorescent chemicals re-emitting light upon light excitation within different ranges of wavelengths. Several types of such fluorophores are commercially available conjugated either to secondary antibodies or for in-house labeling of primary antibodies. They are characterized in terms of brightness, photostability, and environment sensitivity. The commonly used fluorophores include Alexa Fluor, Atto, FITC, and TRITC [6]. Trafficking of endocytosed cargo via pinocytosed plasma membrane progresses via multiple membrane carrier maturation processes. Initially, endocytic membrane carriers fuse intracellularly to form larger organelle sorting stations wherefrom sorting fate are orchestrated [7]. The co-localization of fluorescence from stains of cargo and organelle markers are analyzed in the intracellular space to study where a specific cargo locates during intracellular trafficking. Inclusion of different time points enables a multidimensional mapping of cargo trafficking in BECs. The endo-lysosomal compartment consists of organelles sizing from 100 nm (early endosomes) to more than 1 μm for late endosomes [7]. These sizes are distinguishable by at least one pixel for most high-resolution microscopes. This enables mapping of receptor antibody trafficking to distinct intracellular compartments by 1-pixel two-dimensional (2D) confocal co-localization analysis. The open-source imaging

196

Mikkel R. Holst et al.

analysis software ImageJ offers different methods for analyzing the co-localization of specific cargos and organelle markers. One of them is the analysis plugin for spot detection and co-localization ComDet (https://github.com/ekatrukha/ComDet). The tool is developed for analysis of 2D micrographs and can be used on 3D micrographs containing 2D z-stacks to perform global 3D co-localization analysis. By combining pulse-chase cell trafficking experiments with specific ICC and 1-pixel 2D confocal co-localization analysis, we have developed an assay for mapping of cargo trafficking in BECs. The method allows for identification of surface-expressed receptors of BECs and the degree of co-localization with markers of intracellular compartments at chosen time points using open-source image processing (ImageJ). In this chapter, we map the intracellular trafficking of a chosen transferrin receptor (TfR) antibody from the luminal surface of BECs. The experiments demonstrate a preferred route for the chosen TfR antibody via the early endosome (marked by EEA1) with less trafficking to the late endosome (marked by Rab7) during 20 min internalization. Knowledge of receptor antibody trafficking in BECs is fundamental to understand transcellular trafficking of cargoes from blood to brain and therefore vital to improve receptor-mediated drug delivery to the brain.

2

Materials

2.1 Antibody Endocytosis from the Luminal Plasma Membrane

1. Porcine BECs [8]. 2. The Transwells from Corning (1.12 cm2 surface area, 0.4 μm pores, Polycarbonate Transwell inserts). 3. DMEM with high glucose (4,500 mg/L), L-glutamine, sodium bicarbonate, and sodium pyruvate. Heat-inactivated fetal bovine serum. 4. PBS; Mix 800 ml of distilled water with 8 g of NaCl, 200 mg of KCl, 1.44 g Na2HPO4, and 240 mg KH2PO4. Adjust the buffer to pH 7.4, and add distilled water to 1 l. Aliquot solution and prepare both a RT and 4
C aliquot. 5. Anti-transferrin receptor antibody. 6. Box with ice and cooling elements. 7. Fixation solution: 4% Paraformaldehyde (PFA) in PBS (see Note 1). Store at 4
C.

2.2 Immunofluorescence Staining of Subcellular Markers

1. Permeabilization solution: 0.1% Triton X-100 in PBS (see Note 2). Store at 4
C. 2. Blocking solution: 2% BSA (w/v) in PBS (see Note 3).

Mapping Receptor Antibody Endocytosis and Trafficking in Brain Endothelial Cells

197

Table 1 Antibody overview Primary antibody

Company

Catalogue/lot no.

Dilution Final concentration

Mouse α TfR

Thermo Fisher scientific 13–6800

1:350

1.4 μg/ml

Rabbit α EEA1

Abcam

ab2900

1:100

10 μg/ml

Rabbit α Rab7

Abcam

ab137029

1:100

0.1–0.5 μg/ml

Secondary antibody

Company

Catalogue/lot no.

Dilution Final concentration

Goat α mouse AF488

Thermo Fisher scientific A11001

1:500

4 μg/ml

Donkey α rabbit AF568 Thermo Fisher scientific A10042

1:500

4 μg/ml

3. Washing solution: RT PBS prepared as described in Subheading 2.1. 4. Antibody solutions (see Table 1). 5. Hoechst nuclei staining solution: 1:50,000 in ddH2O. 6. Sterilized 0.15 mm cover slips (number 1.5), rectangular or round. 7. Sterilized microscope glass slides. 8. Fluorescent mounting media. 2.3 Imaging of Samples with Confocal Microscopy

1. Use a high-resolution confocal microscope equipped with a 60–150 objective lens (a 100 objective resolving 160 nm/ pixel was used to obtain data for this chapter), a laser, or LED light source for fluorophore excitation, fluorophore filters, a camera for detection, and software to operate the system.

2.4 Analysis and Quantification of Colocalization Score

1. A standard computer with ImageJ (https://imagej.net/Fiji/ Downloads) and Excel (Microsoft Office) software installed.

3

Methods

3.1 Antibody Endocytosis from the Luminal Plasma Membrane

1. Grow the porcine BECs on two Transwells until they are tight and polarized (see Notes 4 and 5). 2. Prepare 700 μl growth medium (350 μl per top chamber) (see Note 6) with a specific antibody against the receptor target and 1.4 ml (700 μl per bottom chamber) medium without any addition. Use 1 μg/ml as a final antibody concentration for initial experiments, and then adjust to optimal concentration. In many cases, far less is needed to obtain specific stainings. Place the solutions on ice until use.

198

Mikkel R. Holst et al.

3. Place the Transwells on ice in a cold room for 30 min. It is important that the cells are 4
C before the experiment is initiated, since it will block the cellular endocytosis. 4. Remove medium from top and bottom chambers in the Transwells, and add 700 μl precooled medium without antibody (from step 2) to the bottom chamber and 350 μl medium with antibody to the top chamber. As the low temperature has a tendency to loosen the cells from the Transwell filter, this and the following washing steps should be done carefully. In this setup, endocytosis is followed from the luminal side. Endocytosis from the abluminal side can likewise be followed by adding antibody to the lower chamber. 5. Incubate for 90 min on ice in the cold room to allow saturation of antibody binding to the receptor. 6. Wash the top chambers once with ice-cold PBS with the Transwells still placed on ice. This time point represents the so-called 0 time point, e.g., the pool of receptors located on the cell surface labeled with primary antibodies. If one likes to include a surface staining, extra Transwells should be prepared at this step and be directly fixed as described in Subheading 3.2, step 7 without heating. 7. Remove the PBS in Transwells, and add pre-warmed medium (37
C, 700 μl to the top and 1 ml to the bottom). Place the cells in CO2 incubator, and incubate for the desired time points, which in this case is 2 and 20 min. 8. Remove one Transwell at the given time points, wash once with PBS (37
C or room temperature), and fix the cells with 4% (v/v) paraformaldehyde or ice-cold methanol (see Note 7). Following fixation, wash filter membranes once with PBS. 3.2 Immunofluorescence Staining of Subcellular Markers

1. Permeabilize filter membrane seeded with porcine BECs using 0.1% Triton X-100 (v/v) in PBS for 10 min, RT. Wash once in PBS. 2. Block filters for unspecific binding using 2% BSA (w/v) in PBS for 20 min. at RT. 3. Before applying primary antibodies, filters are cut into pieces in order to incubate different pieces with different primary antibodies. Cut the filter according to the number of primary antibodies of interest (see Note 8). 4. Prepare primary antibodies in Eppendorf tubes in concentrations according to the manufacturer’s descriptions (see Note 9). Because the samples have been preincubated with a primary antibody targeting a receptor of interest, the second primary antibody targeting an intracellular marker protein must be generated in another species (see Note 10). The specific descriptions for the applied antibodies are noted in Table 1.

Mapping Receptor Antibody Endocytosis and Trafficking in Brain Endothelial Cells

199

5. Place a piece of Parafilm on a table surface, and add 50 μl droplets of diluted primary antibody onto the film, and place the filter pieces with the cell side down on the top of the droplet. Incubate with primary antibodies for 1 h at RT. 6. For washing, prepare 50 μl droplets of PBS on the Parafilm, and transfer the filter pieces cell side down. Repeat this step three times. 7. Prepare secondary antibodies in Eppendorf tubes according to the manufacturer’s descriptions (see Note 11). Be aware that these are light sensitive and work in the dark at RT. 8. Prepare 50 μl droplets of diluted secondary antibody with Hoechst staining solution (0.125 μg/ml) on the Parafilm, and place the filter pieces with the cell side down on the top of the droplet. Incubate with primary antibodies for 30 h at RT. 9. For washing, prepare 50 μl droplets of PBS on the Parafilm, and transfer the filter pieces to the drops cell side down, and incubate for 5 min, two times. Afterwards, prepare 50 μl droplets of ddH2O, and repeat incubation for 5 min., two times. 10. Finally, mount the filter pieces on microscope glass slides using fluorescent mounting medium and 1.5 mm cover slips washed in ethanol. Leave the slides ON at 4
C or RT in the darkness, and seal with nail polish before imaging. 3.3 Imaging of Samples with Confocal Microscopy

1. Select a confocal scanning or spinning disc microscope with a high magnification objective (e.g., 60–150 magnification) to increase the resolution of each pixel in the obtained micrographs (see Note 12). 2. Test image the samples to make sure that light intensity and exposure time are set, so the signal output of micrographs is without oversaturated pixels (use the saturation indicator in the microscope operating software). 3. Image the Marker (568), Cargo (488), and Hoechst (405) stains (this order of imaging is important for later co-localization analysis) in separate channels with appropriate filters to separate the fluorophore spectra. 4. Image the monolayer of cells from their top to bottom by z-stacks. 5. Save image file with a specific Ex#_568marker_488cargo_Hoechst.

3.4 Analysis of Colocalization Using ImageJ ComDet Spot Detection Plugin

name,

e.g.,

1. Download and follow the instructions for installing the ComDet plugin from: https://imagej.net/Spots_colocalization _(ComDet). 2. Test spot segmentation parameters (perform this step only for a selected amount of files):

200

Mikkel R. Holst et al.

(a) Open a file by dragging the file into the ImageJ main panel. A “Bio-Formats Import Options” window will open; tick on “split channels”; click Ok. (b) A new window for “Bio-Formats Series Options” opens; tick on “Series 1” and click OK. (c) For channel 1 (e.g., marker channel), click image panel> adjust- > Brightness/Contrast. (d) Use “maximum” threshold tab in Brightness/contrast panel to adjust signal so all marker spots can be visually separated from background signal (see Fig. 1). (e) Make sure channel 1 is the active window by clicking on it before running a particle detection test on it: (f) Open Plugins- > ComDet- > Detect particles.

Fig. 1 Signal adjustment before spot overlay. ImageJ control panel, image window with micrograph 20 from the image z-stack file and the Brightness/Contrast panel on the right. The Brightness/Contrast Maximum tab is used to increase the brightness and contrast of the micrograph on the left

Mapping Receptor Antibody Endocytosis and Trafficking in Brain Endothelial Cells

201

Fig. 2 Adjustment of spot segmentation. ImageJ control panel, image window with micrograph 20 from the image z-stack file and the Particle detection control panel for spot segmentation on the right. The settings are displayed in the open fields of the particle display panel with Preview detection ticked on for visualization of segmented spots in yellow shown in the left image window. Compare overlay with image window in Fig. 1

(g) Set “Approximate particle size” to 1, and set “Intensity threshold” to 5, 6, 7... or 10 while ticking “Preview detection” on and off to see if visible marker spots are covered by yellow particles. Make sure not to use too low intensity threshold – it is better to miss some marker spots than to include false-positive spots from low intensity spots, which could be background. It is generally recommended to use 10 as Intensity threshold (see Fig. 2). Once you are satisfied with the particle overlay on the spots of channel 1, note the settings, and click Cancel. (h) Do the same test (step c-g) for channel 2 (e.g., cargo channel), and note the settings. It is again recommended to use approximate particle size 1 and intensity threshold 10. In case other settings are chosen, note the settings and click cancel.

202

Mikkel R. Holst et al.

(i) Close all files. Once a selection of files has been tested for optimal particle overlay parameters (as under 2), the chosen parameters should be used for all files to ensure consistent analysis. 3. Perform the co-localization analysis: (a) Open a file by dragging the file into the ImageJ main panel. A “Bio-Formats Import Options” window will open; tick off “split channels”; click Ok. (b) A new window for “Bio-Formats Series Options” opens; tick on “Series 1,” and click Ok. (c) Open Plugins > ComDet > Detect particles. (d) Tick on “calculate co-localization,” and set “Max distance between colocalized spots” to 1; click Ok. (e) A new window (Detect Particles in channel 1) opens; tick on “Include larger particles?”; set approximate particle size to 1 and intensity threshold to the chosen value found during the test or 10 as recommended in most cases; click OK. (f) Continue this setup for channel 2 (e.g., cargo channel) and channel 3 (Hoechst channel, see Note 13), and complete the analysis by clicking “OK.” (g) A summary window opens up; click the Edit tab > Select all, type Ctrl C, and paste into an Excel sheet. Enter the column names from the ImageJ co-localization analysis summary table above the pasted numbers in the Excel sheet. 3.5 Quantification of Co-localization Score

1. In the Excel sheet, sum the total amount of spots in cargo channel 2 (column D) and total amount of colocalized spots (column E), respectively. Calculate the Co-localization score ¼ (total cargo spots/total colocalized spots)  100% (see Fig. 3). 2. Repeat this procedure until 25–50 cells are analyzed per condition. Repeat the measurements and analysis from a minimum of three independent experiment days. 3. Gather data and present it as suggested in Fig. 4.

4

Notes 1. Be aware that paraformaldehyde (PFA) used in the fixation solution is toxic and must be handled in a fume hood. The 4% PFA fixation solution can be prepared beforehand, aliquoted, and stored at 20
C. When thawed and reaching RT, be sure that the solution is clear without aggregates, and do not refreeze the solution after thawing since this can affect the fixation quality, essential for the staining procedure.

Mapping Receptor Antibody Endocytosis and Trafficking in Brain Endothelial Cells

203

Fig. 3 Calculation of co-localization score. Example of an Excel sheet filled with data from ComDet summary table. Note that column names above the numbers have been filled in according to summary table column names (these names are not included in the copy paste). The co-localization score is given by % total amount of cargo spots colocalized with marker spots of total amount of cargo spots

2. When preparing the permeabilization buffer with 0.1% Triton X-100 in PBS, be aware that the Triton X-100 stock solution is highly viscous and can be difficult to pipette accurately. Use eventually a pipette tip with a larger opening, or weigh out the required amount on a sterile material which is afterwards dropped into the solution and removed when triton has dissolved. 3. For the 2% BSA solution, use a freshly prepared solution. 4. There are many different immortalized and primary types of BECs that each follows different protocols to mature into polarized BEC. We use primary porcine BEC in this example, and details for purification and culturing can be found in Nielsen et al. [8]. 5. In this example, the receptor localization in the endo-lysosomal system is analyzed 2 and 20 min after endocytosis. More time points as well as the zero time point (equal to surface staining) can provide important information. 6. Depending on the antibody and the receptor, a medium without serum may provide better results. There is no need to add antibiotic to the medium since these experiments are relatively fast and development of a bacterial infection is therefore not likely to happen. When investigating luminal endocytosis and intracellular trafficking, one should also be aware that the applied antibody has affinity for an epitope expressed at the luminal extracellular domain of the receptor.

204

A

Mikkel R. Holst et al. Merge Marker channel

EEA1 Cargo channel

Merge Marker channel

Rab7 Cargo channel

Anti-TfR

Hoechst

20 min. pulse-chase anti-TfR Anti-TfR

Hoechst

B

Co-localization score % Anti-TfR co-localized with marker spots of total Anti-TfR spots

20 min. pulse-chase anti-TfR

80 70 60 50 40 30 20 10 0

*

2 min

EEA1 Rab7

20 min

Pulse time

Fig. 4 Presentation of data. (a) Representative micrographs of maximum projected z-stacks from 20 min. Pulse-chase experiments with anti-TfR antibody. Top panels show co-staining of anti-TfR with EEA1 antibody and Hoechst staining to mark the nuclei. Bottom panels show co-staining of anti-TfR with Rab7 antibody and Hoechst staining. Scale bar ¼ 10 μm. (b) Quantification of 1-pixel co-localization (approx. 160 nm/pixel resolution) between cargo and marker. More than 25 cells were analyzed per condition from 3 independent experiment days. Error bars are SEM. Statistics: Two-way Anova, Sidak’s multiple comparisons test P ¼ 0.0125 analyzed in Graphpad Prism

7. Different antibodies require different fixation protocols for optimal results in IF staining. The most common fixatives are paraformaldehyde and methanol. The paraformaldehyde solutions should be prepared freshly for optimal fixation. The methanol should also be fresh (free of water) and 20
C. Both fixatives normally give nice results with 10 min of fixation. If using the methanol fixation protocol, it can be recommended to change the methanol after 1–2 min to remove traces of PBS from the cells. Important considerations and optimization for proper fixation include penetration rate, temperature,

Mapping Receptor Antibody Endocytosis and Trafficking in Brain Endothelial Cells

205

and the length of fixation. Cooling cells and fixatives can affect the fixation and require longer incubation times for proper fixation. 8. Cut filter membranes into pieces by use of a sharp scalpel blade. We recommend each filter to be divided into maximum four pieces. Change blade regularly to obtain sharp edges since flossy edges will affect the focus area when imaging the samples. 9. Antibody solutions are prepared in Eppendorf tubes. Before pipetting the recommended concentration of antibodies from stocks, quickly spin these down in order to avoid antibody aggregates causing background signal on the samples. 10. When selecting the primary antibodies for receptors and intracellular proteins of interest in the assay, be aware to select primary antibodies raised in different host species and secondary antibodies with different fluorophore excitation wavelengths. For example, incubation with a mouse IgG primary antibody can be used in combination with a rabbit IgG primary antibody followed by incubation with an Alexa-488 goat antimouse IgG secondary antibody and an Alexa 568 goat anti rabbit IgG secondary antibody. 11. Preparation of primary antibodies according to the manufacturer’s description often involves an optimization of the antibody concentration appropriate to the specific experimental setup, including testing of dilutions of 1:100–1:1000. 12. The chosen objective lens affects the image resolution: 60, 100, and 150 objectives will result in approximately 266 nm/pixel, 160 nm/pixel, and 106 nm/pixel resolution, respectively. 13. The analysis for channel 3 will not be used but is run to bypass time-consuming removal of the Hoechst channel from the raw imaging file before co-localization analysis. References 1. Mercer J, Greber UF (2013) Virus interactions with endocytic pathways in macrophages and dendritic cells. Trends Microbiol 21 (8):380–388 2. Toth AE, Holst MR, Nielsen MS (2020) Vesicular transport machinery in brain endothelial cells: what we know and what we Don’t. Curr Pharm Des 26:1405 3. Tomoda H, Kishimoto Y, Lee YC (1989) Temperature effect on endocytosis and exocytosis by rabbit alveolar macrophages. J Biol Chem 264 (26):15445–15450 4. Bordeaux J et al (2010) Antibody validation. BioTechniques 48(3):197–209

5. Howat WJ, Wilson BA (2014) Tissue fixation and the effect of molecular fixatives on downstream staining procedures. Methods 70 (1):12–19 6. Panchuk-Voloshina N et al (1999) Alexa dyes, a series of new fluorescent dyes that yield exceptionally bright, photostable conjugates. J Histochem Cytochem 47(9):1179–1188 7. Huotari J, Helenius A (2011) Endosome maturation. EMBO J 30(17):3481–3500 8. Nielsen SSE et al (2017) Improved method for the establishment of an in vitro blood-brain barrier model based on porcine brain endothelial cells. J Vis Exp 127

Methods in Molecular Biology (2021) 2367: 207–213 DOI 10.1007/7651_2021_390 © Springer Science+Business Media, LLC 2021 Published online: 09 April 2021

An In Vitro Assay to Monitor Sertoli Cell Blood-Testis Barrier (BTB) Integrity Siwen Wu, Lingling Wang, Elizabeth I. Tang, Junlu Wang, and C. Yan Cheng Abstract In this chapter, we detail a reliable, effective, and easy to perform assay to monitor the Sertoli cell blood–testis barrier (BTB) integrity. While the BTB in the testis is composed of the tight junction (TJ) barrier and basal ES (ectoplasmic specialization, a testis-specific actin-rich adherens junction (AJ) type), this method is applicable to all other blood–tissue barrier in vitro, including endothelial TJ-barrier of the blood–brain barrier (BBB). Furthermore, this method does not require expensive set up, and can be rapidly performed by any standard biochemistry/cell biology/molecular biology laboratory. The basic idea is built on the concept that a functional blood–tissue barrier, such as the BTB conferred by Sertoli cells in the testis, is capable of blocking the diffusion of a small membrane impermeable biotin (e.g., EZ-Link Sulfo-NHS-LCbiotin, Mr. 556.59) across the barrier. However, when this barrier is compromised, such as following treatment with a toxicant or knockdown of a relevant gene necessary to confer the TJ-barrier function, the biotin will permeate the barrier, reaching the Sertoli cell cytosol. Biotin can be subsequently visualized by using streptavidin conjugated to a fluorescence tag such as Alexa Fluor 488 (green fluorescence) which can be easily visualized by a standard fluorescence microscope. Key words Blood–barrier, Testis, BTB integrity assay, Sertoli cells, Spermatogenesis, Testis

1

Introduction The function of a blood–tissue barrier in the mammalian body, such as the blood–testis barrier (BTB) in the testis, the blood–brain barrier (BBB) in the brain, or the blood–retina barrier (BRB) in the eye, is to create a unique microenvironment to support important cellular function in an organ [1–5]. For instance, the BTB created by Sertoli cell TJ-barrier and supported by basal ES at the base of the seminiferous tubule in the testis versus the BBB created by endothelial TJ-barrier of the microvessel (or capillary) and supported by pericytes in the brain. These blood–tissue barriers provide the necessary transcellular (across cells, the fence function) and paracellular (between cells, the gate function) diffusion of water, electrolytes, minerals, and biomolecules. In the testis, the BTB also divides the seminiferous epithelium into the basal and the apical (adluminal) compartments so that meiosis I/II and post-meiotic

207

208

Siwen Wu et al.

development of haploid spermatids take place in the apical compartment, behind the BTB. Studies have shown that a delay of BTB assembly in rats by treatment of neonatal rats with diethylstilbestrol (a synthetic estrogen) also delays the occurrence of meiosis and the appearance of haploid germ cells, impeding spermatogenesis [6], consistent with treatment of neonatal rats with a goitrogen (e.g., 6-propyl-2-thiouracil, PTU) [7]. Taken collectively, these studies illustrate the importance of BTB establishment by ~day 17–19 dpp (day postpartum) in Sprague–Dawley rats to support the onset of spermatocyte maturation to initiate meiosis I/II [8]. A functional assay to monitor BTB integrity in vivo is available [9]. On the other hand, a reliable physiological assay to monitor the Sertoli cell TJ-barrier in vitro by quantifying the resistance of a functional BTB to the passage of current across the Sertoli cell tight junctions is also available [10]. This latter method is tedious as it also requires expensive equipment (e.g., a Millipore ERS meter with two silver-silver chloride electrodes) and more sophisticated skill [10, 11]. Herein, we provide a detailed protocol for assessing the Sertoli cell BTB in vitro based on the use of a membrane impermeable biotin [12]. This method is reliable, easy to perform without the use of sophisticated equipment and time consuming training, and can be easily performed by any lab personnel in a regular life science laboratory.

2

Materials 1. Ten 20-day-old male Sprague–Dawley pups (Charles River Laboratories, Kingston, New York). 2. 12-Well Culture Plate coated with BD Matrigel™ Basement Membrane Matrix (Cat #: 354234) diluted at 1:7 with DMEM/F-12 and placed inside a water jacketed carbon dioxide (CO2) incubator with 95% air/5% CO2 at 35
C. 3. Pipettes. 4. EZ-Link™ Sulfo-NHS-LC-Biotin (Mr 556.59, a cell membrane impermeable biotinylation reagent, ThermoFisher, Cat #: 21335). 5. Dulbecco’s Modified Eagle’s Medium/Nutrient Mixture F-12 Ham (Millipore Sigma, Cat # D8437). 6. Phosphate-buffered saline (PBS): 10 mM NaH2PO4, 0.15 M NaCl, pH 7.4 at 22
C. 7. Phosphate-buffered saline (PBS)/CM: 10 mM NaH2PO4, 0.15 M NaCl, 1 mM CaCl2, pH 8.0 at 22
C. 8. Labeling buffer: 10 mg/ml EZ-Link™ Sulfo-NHS-LC-Biotin (ThermoFisher Cat #:21335) in PBS/CM pH 8.0 containing 1 mM CaCl2.

In Vitro BTB Integrity Assay

209

9. 70% Ethanol (vol/vol, in MilliQ water). 10. 10 mL, Polystyrene Serological pipettes. 11. 4% PFA (paraformaldehyde) in PBS. 12. Streptavidin conjugated with Alexa Fluor 488 (green fluorescence, ThermoFisher, Cat #: S11223) (streptavidin conjugated with other types of fluorophore, such as Cy3 for red fluorescence, can also be used based on specific experimental needs). 13. ProLong® Gold Antifade Mountants reagent (ThermoFisher, Cat #: P36930). 14. 40 ,6-diamidino-2-phenylindole (DAPI, Life Technologies Corporation, Cat #: P36935). 15. Microscope slides (Thomas Scientific, Cat #: 6686S50). 16. Standard Fluorescence Microscope (such as Nikon Eclipse 90i equipped with Nikon Ds-Qi1Mc and DsFi1 digital camera) or confocal microscope (such as Zeiss LSM 880 NLO laser scanning confocal and multiphoton microscope).

3 3.1

Methods In Preparation

1. Isolate Sertoli cells from ten 20-day-old male Sprague–Dawley pups as detailed elsewhere [10] and plate them in six 12-well culture plates at a low cell density (0.02  106 cells/cm2) (see Note 1) and culture the cells in 3-mL of DMEM/F-12 for 4–5 days to allow the establishment of a functional permeability barrier as described [10]. Media are to be replaced every 48 h and supplemented with growth factors [10]. 2. On the day of experiment to monitor Sertoli cell BTB integrity, clean the hood with 70% ethanol before experiment and warm 100 mL DMEM/F-12 to 35
C in a shaking water bath. 3. Immediately before use, equilibrate all media to room temperature. 4. Freshly prepare a stock of 10 mg/ml EZ-Link™ Sulfo-NHSLC-Biotin in PBS/CM pH 8.0 (10 mM sodium phosphate, 0.15 M NaCl) containing 1 mM CaCl2 as labeling buffer (biotin reagent is freshly prepared immediately before use, within 1–2 h before its use; do not prepare stock solution for storage) (see Note 2).

3.2

Experiment

1. Wash Sertoli cells three times by rinsing cells with ice-cold PBS (pH 7.4) to remove amine-containing media and proteins from the cells. 2. Add 20 μL of the biotin reagent per mL DMEM/F-12 which served as the Biotinylation Reaction Mixture. This reaction

210

Siwen Wu et al.

mixture should be prepared shortly prior to its use from the stock solution in Subheading 3.1, step 4 (see Note 2). 3. Add 1 mL of Biotinylation Reaction Mixture to each well of the 12-well plate. 4. Incubate reaction mixture at room temperature for 30 min. 5. Wash cells three times with PBS + 100 mM glycine to quench the biotinylation reaction, and gently remove excess biotin reagent and byproducts by aspiration. 6. Fix cells in ice-cold methanol at room temperature for 5 min (or cells can be fixed in 4% PFA (paraformaldehyde) in PBS for 5 min, wash thrice in PBS at room temperature, and then permeabilized in 0.1% Triton X-100 for 10 min, this step is not necessary using ice-cold methanol to fix Sertoli cells). 7. Wash with PBS thrice at room temperature. Incubate with blocking solution (1% BSA in PBS, wt/vol) for 30 min at room temperature. 8. Remove blocking solution by aspiration. Add ~50 μL of Streptavidin conjugated with Alexa Fluor 488 (green fluorescence) using 1:200 dilution (to visualize biotin) containing 40 ,6-diamidino-2-phenylindole (DAPI, to visualize Sertoli cell nuclei). Incubate for 1 h at room temperature (see Note 3). 9. Wash with PBS thrice at room temperature. Dry coverslips in dark. Mount onto microscope slides with ProLong® Gold antifade reagent (see Note 3). 3.3 Image Requisition and Data Analysis

1. Acquire images using a standard fluorescence microscope or confocal microscope. Typical fluorescent images are shown in Fig. 1, which were obtained using the Nikon Eclipse 90i Fluorescence Microscope system. These findings illustrate the intact BTB between normal Sertoli cells (Normal/Ctrl) is capable of blocking the diffusion of membrane impermeable biotin from crossing the barrier to enter Sertoli cells. However, following transfection of Sertoli cell epithelium with KIF15-specific siRNA duplexes to knockdown KIF15 in which its expression has been reduced by ~70% as recently reported [12], and the Sertoli cell tight junction barrier has also been compromised based on a physiological assay of monitoring BTB integrity based on TER (transepithelial electrical resistance) measurement, biotin effectively permeates the Sertoli cell BTB (Fig. 1). Yet, Sertoli cell epithelium transfected with non-targeting siRNA duplexes is able to block the diffusion of biotin across the cell tight junction barrier (Fig. 1), similar to the control Sertoli cells (Normal/Ctrl) without any treatment. 2. The following steps are used in experiments wherein the findings noted in Fig. 1 are intended to be semiquantitatively

In Vitro BTB Integrity Assay

Ctrl RNAi

KIF15 RNAi

Biotin/DAPI

Normal/Ctrl

211

Fluoresence at the cell-cell interface (Arbitrary unit)

Ctrl RNAi KIF15 RNAi 5

**

4 3 2 1 0

Fig. 1 A study to assess Sertoli cell tight junction (TJ) barrier function that mimics the Sertoli cell BTB in vivo. Sertoli cell density was maintained at 0.02  106 cells/cm2 in the experiment reported here. Representative images illustrating the result of an in vitro Sertoli cell BTB integrity assay wherein a functional TJ-barrier is capable of blocking biotin (green fluorescence, EZ-Link sulfo-NHS-LC-biotin, a membrane impermeable biotin, visualized by Alexa Fluor 488-streptavidin) from entering the Sertoli cell cytosol as noted in control Sertoli cell epithelium (left panel). In Sertoli cells transfected with KIF15 siRNA duplexes (100 nM), beginning on day 2 and cells were washed on day 3 to remove the transfection medium [12]. Thereafter, cells were incubated with fresh DMEM/F12 and cultures were terminated on day 4 for the BTB integrity assay (see Protocols above), the BTB was perturbed (right panel). It is noted that biotin freely permeated into the Sertoli cell cytosol, consistent with findings recently reported wherein the BTB integrity was monitored by other parameters in addition to this in vitro assay [12]. However, Sertoli cells transfected with nontargeting siRNA duplexes which served as the negative control, the Sertoli cell TJ-barrier was not affected and biotin was blocked from entering the Sertoli cell cytosol but remained at the Sertoli cell–cell interface (middle panel). Relative distribution of biotin at the Sertoli cell–cell interface in controls (Ctrl RNAi, transfected with negative non-targeting siRNA duplexes) was annotated by a white bracket vs. a yellow bracket in KIF15-silenced cells wherein cells were transfected with KIF15-specific siRNA duplexes [12]. Red siGLO (red fluorescence) noted in middle and right panel illustrate successful transfection, and no transfection was performed in the control on the left panel. Bar graph on the lower panel is the composite semiquantitative data, with each bar representing a mean  SD of n ¼ 3 experiments. Scale bar, 20 μm, which applies to other panels

212

Siwen Wu et al.

presented to quantify the extent of BTB disruption. Investigators can measure the distance traveled by biotin in the Sertoli cells in the test group (e.g., cells transfected with KIF15 siRNA duplexes) (DTest–biotin) (see yellow brackets) and the distance traveled by biotin in the normal Sertoli cells (or cells transfected with nontargeting control siRNA duplexes) (DCtrl–biotin) (see white brackets). For an oval-shaped Sertoli cells, the distance is the average of the shortest and the longest distance traveled by biotin in the Sertoli cytosol. The extent of the BTB damage (E) can be expressed in percentage (n ¼ 3 experiments, a total of ~200 randomly selected Sertoli cells by scoring about 70 Sertoli cells in each experiment) as: E ¼ ½DTestbiotin =DCtrlbiotin   100% 3. At least 200 Sertoli cells are randomly selected and quantified from a total of n ¼ 3 experiments. 4. Calculate the E for each experimental group, and compare each group using appropriate statistical method. 3.4

4

Conclusion

This simple in vitro assay to monitor Sertoli cell BTB integrity has been used by several investigators in our research group with satisfactory results as noted in Fig. 1. Furthermore, this technique has been used to present findings semiquantitatively following RNAi of a microtubule-dependent plus (+) end directed motor protein KIF15 published in a recent report from our laboratory [12], and also summarized in Fig. 1. We also anticipate that this technique is applicable to other epithelial/endothelial cells cultured in vitro and to assess if they are also capable of establishing a blood–tissue barrier, such as endothelial cells that mimic the BBB in the brain.

Notes 1. For studies using Sertoli cells as the study model to examine BTB as a blood–tissue barrier to monitor barrier integrity in vitro, we suggest a cell density of 0.02–0.03  106 cells/ cm2 so that cells can be evenly spaced and the intercellular junctions can be easily visualized by fluorescence microcopy. However, for other blood–tissue barriers, such as the use of endothelial cells to examine BBB integrity, investigators can consider adjust the optimal cell density in pilot experiments at the range of 0.01–0.05  106 cells/cm2 to obtain the best results. 2. The biotinylation stock solution of 10 mg/ml EZ-Link™ Sulfo-NHS-LC-Biotin in PBS/CM pH 8.0 (10 mM sodium phosphate, 0.15 M NaCl) containing 1 mM CaCl2 should be

In Vitro BTB Integrity Assay

213

freshly prepared within 1–2 h prior to its use to serve as the Biotinylation Reaction Mixture. 3. Following addition of streptavidin conjugated with Alexa Fluor 488 (green fluorescence) containing 40 ,6-diamidino-2-phenylindole (DAPI, to visualize Sertoli cell nuclei), it is necessary to acquire images between treatment and control groups to assess Sertoli cell BTB integrity within 12 h to avoid fading of the fluorescence signals. Furthermore, images between control and treatment groups should be acquired within the same experimental session to avoid inter-experimental variations.

Acknowledgments This work was supported by a grant from The Wenzhou Medical University (to C.Y.C.) References 1. Cheng CY, Mruk DD (2012) The blood-testis barrier and its implication in male contraception. Pharmacol Rev 64:16–64 2. Easton AS (2012) Regulation of permeability across the blood-brain barrier. Adv Exp Med Biol 763:1–19 3. Mruk DD, Cheng CY (2004) Sertoli-Sertoli and Sertoli-germ cell interactions and their significance in germ cell movement in the seminiferous epithelium during spermatogenesis. Endocr Rev 25:747–806 4. Stanton PG (2016) Regulation of the bloodtestis barrier. Semin Cell Dev Biol 59:166–173. https://doi.org/10.1016/j.semcdb.2016.06. 018 5. Campbell M, Humphries P (2012) The bloodretina barrier: tight junctions and barrier modulation. Adv Exp Med Biol 763:70–84 6. Toyama Y, Maekawa M, Yuasa S (2003) Ectoplasmic specializations in the Sertoli cells: new vistas based on genetic defects and testicular toxicology. Anat Sci Int 78:1–16 7. De Franca L, Hess R, Cooke P, Russell L (1995) Neonatal hypothyroidism causes delayed Sertoli cell maturation in rats treated with propylthiouracil: evidence that the Sertoli

cell controls testis growth. Anat Rec 242:57–69 8. Mok KW, Mruk DD, Lee WM, Cheng CY (2011) A study to assess the assembly of a functional blood-testis barrier in developing rat testes. Spermatogenesis 1:270–280 9. Chen H, Lui WY, Mruk DD, Xiao X, Ge R, Lian Q et al (2018) Monitoring the integrity of the blood-testis barrier (BTB): an in vivo assay. Methods Mol Biol 1748:245–252. https:// doi.org/10.1007/978-1-4939-7698-0_17 10. Mruk DD, Cheng CY (2011) An in vitro system to study Sertoli cell blood-testis barrier dynamics. Methods Mol Biol 763:237–252 11. Grima J, Wong CC, Zhu LJ, Zong SD, Cheng CY (1998) Testin secreted by Sertoli cells is associated with the cell surface, and its expression correlates with the disruption of Sertoligerm cell junctions but not the inter-Sertoli tight junction. J Biol Chem 273:21040–21053 12. Wu S, Lv L, Li L, Wang L, Mao B, Li J et al (2021) KIF15 supports spermatogenesis via its effects on Sertoli cell microtubule, actin, vimentin, and septin cytoskeletons. Endocrinology 162(4):bqab010. https://doi.org/10. 1210/endocr/bqab010

Methods in Molecular Biology (2021) 2367: 215–233 DOI 10.1007/7651_2020_324 © Springer Science+Business Media New York 2020 Published online: 19 September 2020

Ussing Chamber Methods to Study the Esophageal Epithelial Barrier Solange M. Abdulnour-Nakhoul and Nazih L. Nakhoul Abstract The Ussing chamber was developed in 1949 by Hans Ussing and quickly became a powerful tool to study ion and solute transport in epithelia. The chamber has two compartments strictly separating the apical and basolateral sides of the tissue under study. The two sides of the tissue are connected via electrodes to a modified electrometer/pulse generator that allows measurement of electrical parameters, namely, transepithelial voltage, current, and resistance. Simultaneously, permeability of the tissue to specific solutes or markers can be monitored by using tracers or isotopes to measure transport from one side of the tissue to the other. In this chapter, we will describe the use of the Ussing chamber to study the barrier properties of the mouse esophageal epithelium. We will also briefly describe the use of the modified Ussing chamber to simultaneously study transepithelial and cellular electrophysiology in the rabbit esophageal epithelium. Lastly, we will cover the use of the Ussing chamber to study bicarbonate secretion in the pig esophagus. These examples highlight the versatility of the Ussing chamber technique in investigating the physiology and pathophysiology of epithelia including human biopsies. Key words Transepithelial, Voltage, Resistance, Short-circuit current, Bicarbonate, pH stat, Stratified squamous, Epithelium, Microelectrodes, Agar bridges, Ion transport, Permeability, Voltage clamp

1

Introduction Epithelial cells are held together by intercellular junctions to form sheets of tissue that cover the external surface of the body and line the surface of the gastrointestinal, respiratory, excretory, and reproductive tracts. Epithelial cells are polarized as they have an apical (facing outside or luminal side) and a basolateral membrane (facing blood side). They play important roles in protecting the underlying tissues and in maintaining a stable internal “milieu.” They also have a major role in absorption and secretion. Epithelial transport of ions and solutes could be primary or secondary active [1], or it could be passive through pores and channels [2]. The Ussing chamber has been instrumental to study ion transport in a variety of epithelial tissues including frog skin, toad urinary bladder [3], airways [4], intestine [5], gallbladder [6, 7], stomach [8], and esophagus [9, 10–12].

215

216

Solange M. Abdulnour-Nakhoul and Nazih L. Nakhoul

Fig. 1 Hematoxylin & eosin staining of mouse (a) and pig (b) esophageal tissue sections. Submucosal glands (SMG) are present only in pig tissue. SB stratum basalis, SS stratum spinosum, SG stratum granulosum, SC stratum corneum. Bar ¼ 100 μm in a and 1 mm in b

The esophagus is lined with stratified squamous epithelium consisting of multiple cell layers that cover and protect underlying structures from chemical injuries due to ingested food and periodic reflux of acidic stomach contents. According to the KoefoedJohnsen and Ussing, two-membrane model [13] Na+ enters the cell passively across the luminal membrane through Na+ channels and exits across the basolateral membrane via the action of the Na+K+ pump [9]. The basal cells (stratum basalis) of the esophageal epithelium continuously multiply and form new layers that migrate to the surface. As the cells differentiate and advance toward the lumen, they lose their nuclei and eventually desquamate. In rodents, unlike humans, the uppermost layer is keratinized (Fig. 1a). The rate of regeneration of the whole epithelium in the mouse is about 7 days [14, 15], and in humans, it is approximately 11 days [16]. The integrity of the barrier is maintained through this process. The overall barrier function of the epithelium is dependent on three elements: The first one, the paracellular pathway, depends on the composition of the intercellular spaces and the junctions between the cells. The second one depends on the composition of the cell membranes and transcellular ion transport. The third component depends on pre-epithelial buffering. Factors disrupting these elements could lead to esophageal disease including GERD, Barrett, and carcinogenesis [17–20]. The cellular pathways for ion transport are responsible for the maintenance of intracellular homeostasis. This is particularly important in the esophageal epithelium which is constantly exposed to various noxious luminal agents ranging from high acidity, especially during episodes of gastroesophageal reflux, to hypertonicity of ingested food and beverages. Defining the specific pathways contributing to ion transport across the cell membranes is crucial to understand the contribution of cellular transport for

Methods to Study the Esophageal Epithelial Barrier

217

maintenance of the barrier function of the tissue. Apical or basal cells of the epithelium can be impaled with microelectrodes. Microelectrodes are utilized to measure intracellular Na+ or Cl activity, intracellular pH, and membrane potential of esophageal cells within the intact epithelium. It is a powerful tool to understand ion transport at the cellular level in the tissue [21, 22]. In human esophagus, pre-epithelial buffering is a function of the esophageal submucosal glands that contribute significantly to epithelial defense by secreting HCO3 (and mucin) in amounts sufficient to neutralize residual volumes of acid left in the esophagus after bolus clearance [23]. Quantitatively, this HCO3 secretion can approach the HCO3 output of salivary glands at rest [24, 25]. During sleep, when esophageal clearance mechanisms such as upright position, swallow-induced peristalsis, and salivary secretion are inoperative, the secretions from the glands become a unique contributor to acid clearance and pre-epithelial protection. This gradient was evident in tissues from the opossum, which contain SMG, but was absent in tissues from the rabbit or rodents, which do not contain SMG. In the opossum this gradient was significantly enhanced (threefold) by serosal application of carbachol (CCh), a cholinergic agonist, which resulted in a surface pH approximately 2 pH units higher than the luminal pH of 3.5 [26]. Considering that the median luminal pH value in the human esophagus is around 4 [27], this protective mechanism is significant. Among species that have esophageal submucosal glands are humans, opossums, dogs, and pigs [28]. In the pig esophagus, the submucosal glands are located in the orad part of the tissue (Fig. 1b). Alkaline secretion in the pig esophagus was found to be dominated by HCO3 secretion [29–31]. We will describe the use of the Ussing chamber to (1) study the barrier properties of the mouse esophageal epithelium as it relates to intercellular properties; (2) study cellular ion transport in basal and luminal rabbit esophageal cells; (3) and measure bicarbonate secretion in the pig esophagus.

2

Materials

2.1

Solutions

Stirrer; pH meter; calibrated pH electrode; 1 N NaOH; HEPES free acid (238.3 g/mol); NaCl (58.44 g/mol); K2HPO4 (174.2 g/ mol); KH2PO4 (136.09 g/mol); MgCl2·6H2O (203.31 g/mol); CaCl2 anhydrous (110.99 g/mol); D-glucose (180.2 g/mol); NaHCO3 (84.01 g/mol); oxygen gas cylinder fitted with regulator for bubbling; CO2 gas cylinder fitted with regulator for bubbling; deionized water

2.2

KCl Agar Bridges

Pipettor (200 μL); modified gel loading pipette tips (Physiologic Instruments P2023-20); KCl; agar powder; food and drug coloring green (FDC green #3); double boiler (500 ml beaker, 100 ml beaker); glass beads (boilers); 10–12-in.-straight forceps

218

Solange M. Abdulnour-Nakhoul and Nazih L. Nakhoul

2.3 Ussing Chambers

Physiologic instruments VCC M4 multichannel voltage/current clamp; water-jacketed stand; chambers and sliders (for mouse esophagus P2407 2.0 mm diameter aperture for area ¼ 0.031 cm2); 8 sintered Ag/AgCl pellet electrodes for voltage sensing (black); 8 Ag wire electrodes for current passing (white); agar-filled bridges; Ringer’s solution; circulating water bath connected to the stand to warm the chambers; water bath

2.4 Mouse Tissue Dissection and Mounting

Stereoscope; freshly excised mouse esophagus; ice-cold HEPES Ringer; scissors; Iris forceps; surgical spring scissors; Dumont #5 forceps; petri dishes; ice

2.5

Circuit Analysis

Data acquisition system (DATAQ Instruments, DI 720); computer loaded with Acquire and Analyze program

2.6

Fluorescein Flux

Fluorescence microplate reader (excitation wavelength 485 nm, emission wavelength 528 nm), fluorescein isothiocyanate dextran (FITC-Dextran, MW 4 kDa), 96-well plates (clear, flat bottom), pipettors, vortex mixer, aluminum foil, HEPES Ringer at pH 7.5 and at pH 1.6

2.7

Low pH Ringer

Use the components described in Subheading 2.1 above with the following modifications: omit HEPES free acid, adjust the amount of NaCl to 8.18 g/l, and titrate the pH to 1.6 with 5 N and 1 N HCl acid. The solution can be refrigerated and kept at 4  C for up to a month.

2.8 Microelectrode Measurements in Esophageal Tissues Mounted in a Modified Ussing Chamber

Vibration isolation table (Newport); Faraday cage; Ultraprecise Manual Control Micromanipulator (Warner Instruments) for microelectrode placement; (MM-33) Joystick Manual Micromanipulators for placement of reference electrodes; 6-way and 4-way rotary and slider valves (Rheodyne, 5012 Six-way and 5042 FourWay Teflon Rotary Valves, 1.5 mm bore) for quick switching of perfusion solutions with minimal dead-space; Clippard pneumatic valve; nitrogen tank; VCC 600 Voltage-Current Clamp (Physiologic Instrument); high impedance electrometer FD223 (WPI); borosilicate glass tubing 1.2 OD, 0.6 ID (Warner Instruments); electric shrinkable tubing; water bath to warm the solutions to 37  C; freshly excised rabbit esophagus; computerized data acquisition system (Data-Trax WPI, FL)

2.9 Bicarbonate Secretion Measurement in the Ussing Chamber

Radiometer pH stat system fitted with a pH mini-electrode PHC 4000; Ussing chambers set-up as described in Subheadings 2.2 and 2.3 above; pig esophagus; P2315 sliders for the Ussing chambers (12.7 mm diameter; A ¼ 1.27 cm2); bicarbonate Ringer’s solution; saline NaCl 0.9 g/100 ml of deionized water, carbachol (Sigma Aldrich); HCl 0.01 N

Methods to Study the Esophageal Epithelial Barrier

3 3.1

219

Methods 1. Calibrate a pH electrode using the appropriate buffers as required for the pH meter in use. To prepare 1 l of mammalian HEPES Ringer’s solution, add 700 ml of deionized water in a 1-liter beaker. Add a magnetic stir bar and place on a stirrer. While stirring, add in order the compounds as described in Table 1.

Solutions

2. After addition of all solutes, place the calibrated pH electrode in the solution to measure the pH. Slowly titrate the Ringer’s solution with 1 N NaOH until pH of 7.4 is reached (approximately 12.5 mM of HEPES free acid will be titrated as a Na+ salt at pH 7.4, (pKa ¼ 7.4 at 25  C and 7.3 at 37  C). 3. Measure the osmolality of the solution which should be ~290 mOsm/kg H2O. The final composition of Ringer’s solutions will be in mM: Na+, 145; Cl, 137.8; K+, 5.2; HEPES, 25; Ca2+, 1.2; Mg2+, 1.2; HPO42, 2.4; H2PO4, 0.4; glucose, 10, (osmolality 290 mOsm/kg H2O, pH 7.4, gassed with 100% O2). The solution can be refrigerated and kept at 4  C for up to 10 days (see Notes 1, 2, 3, and 4). 4. To prepare bicarbonate Ringer, add NaHCO3 to a beaker containing 300 ml of deionized water. Bubble with 5% CO2 (balance O2) for 30 min. 5. Add all the other components to another beaker containing 600 ml of deionized water. Adjust the pH to 7.4 if needed with 0.1 N NaOH or 0.1 N HCl. Add the bubbled bicarbonate Table 1 Composition of solutions Bicarbonate Ringer (g/l)

Bicarbonate Ringer (mmol/l)

0

0

7.01

Na+, 120; Cl, 120

K+, 5.2; HPO42, 2.4 H2PO4, 0.4

0.418 0.054

K+, 5.2; HPO42, 2.4 H2PO4, 0.4

MgCl2.6H2O 0.243

Mg2+, 1.2; Cl, 2.4

0.243

Mg2+, 1.2; Cl, 2.4

CaCl2 anhydrous

0.133

Ca2+, 1.2; Cl, 2.4

0.133

Ca2+, 1.2; Cl, 2.4

D-glucose

0.9

D-glucose, 10

0.9

D-glucose, 10

NaHCO3

0

0

2.10

Na+, 25; HCO3, 25

Compound

HEPES Ringer HEPES Ringer (g/l) (mmol/l)

HEPES (free acid) NaCl

5.957 7.77

K2HPO4 KH2PO4

0.418 0.054

Na+, 133; Cl, 133 HEPES, 25

220

Solange M. Abdulnour-Nakhoul and Nazih L. Nakhoul

solution to the mixture. Adjust the volume to 1 l. Keep bubbling for an additional 10 min. pH will be 7.4 during bubbling with CO2. Important: pH should not be adjusted after NaHCO3 addition because it results in titration of HCO3 and its loss as CO2 (see Notes 1, 2, 3, and 5). 3.2

KCl Agar Bridges

1. Prepare 500 ml of 3 M KCl by adding 111.83 g of KCl (molar mass: 74.5513 g/mol) in 500 ml deionized water. Stir gently until dissolved. 2. To make the double boiler: place enough glass beads in the 500 ml beaker to a height of ~2.5 cm (~1 in), and add about 200 ml of distilled water. Add 50 ml of 3 M KCl to the 100 ml beaker, and add 1.5 g of agar (3%) and 0.05 g of FDC green (0.1%) and a small stir bar. Place the 100 ml beaker over the beads in the 500 ml beaker; this will be the inner part of the double boiler. Place on a hot plate stirrer, and bring to a gentle boil while gently stirring. Boil the agar for about 10 min; then while the solution is still on low heat, fit each pipette tip to the pipettor, and draw about 50 μl of the boiling agar mixture into the tip, and, without taking the pipette tip out of the solution, release the tip from the pipette using the long forceps. The green dye helps to visualize the agar column to make sure it is continuous and not interrupted by air bubble. Fill about 25 tips, place the agar-filled tips on a paper towel, and let them set for about 30 min. 3. Prepare a storing solution for the tips by adding 0.1 g of FDC green to 100 ml of 3 M KCl (0.1%). Backfill each pipette with the storing solution using a 1 ml syringe fitted with a luer lock needle. Place the agar-filled pipettes in the 3 M KCl storing solution. Cover and refrigerate. The filled electrodes can be stored up to a month in the fridge.

3.3 Ussing Chambers Set-Up

1. Turn equipment on in the following order: the electrometer, data acquisition box, and computer. 2. Place the chambers in the stand. Insert the sliders (without tissues) between the chambers and tighten the clamps. 3. Take the agar-filled electrodes out of the 3 M KCl storage solution using plastic forceps, and rinse the tips by dipping it in a beaker of deionized water. Make sure the agar reaches the tip. If not, make a diagonal cut using a sharp blade. Connect eight agar-filled electrodes to the Ag/AgCl wire electrodes (white) for current passing, making sure that no air bubbles interrupt the liquid column. Insert the agar tip firmly in in the chamber opening farthest from the slider on all chambers. Connect eight more agar tips to the sintered Ag/AgCl electrodes for voltage measurement, and insert in the holes closest to

Methods to Study the Esophageal Epithelial Barrier

221

the slider. V1 indicates the basolateral side of the tissue and V2 indicates the luminal side. Connect the color-coded pins from the “single channel input module” (DM_MC6) to each electrode. Fill each chamber with 5 ml of warm HEPES Ringer. 4. Open the readout on each panel of the voltmeter to read the baseline voltage which is the asymmetry potential difference between the sets of electrodes and should be no more than few millivolts (90% IPA for 15 min in Form Wash station. 4. Cure in Form cure for 15 min at 60  C in Form Cure station. 5. Remove any support material with the use of flush cutters and a Dremel. 6. Clean the threads using taps and dies. The stretching apparatus has five parts: two fasteners on the steady edge, one peg, one pulling clamp, and the main container. A starting point is marked in a side of the main container where the length from the steady edge to the starting point is 5 cm. This device is designed permitting the tensile strain equation shown below (Eq. 1), where ΔL is the change in length and L0 is the starting length [40]. Then, the ratio between ΔL and L0 is converted to a percentage to obtain the strain percentage (Eq. 2). Hence, at 0% strain the length of the membrane or L0 is 5 cm. Tensile strain ¼

ΔL L0

  ΔL  100% Tensile strain percentage% ¼ L0

ð1Þ ð2Þ

Two fasteners are designed to keep the membrane stable and locked at the steady edge. Thus, the membrane does not move at this end. The uniaxial strain is applied by pulling the clamp via rotating the peg. The peg is marked with numbers that represent

A Simple Stretch Assay to Study Strain on Epithelial Cells

239

Fig. 1 The stretching device. (a) The fully assembled device is comprised of five parts: (b) two fasteners, the peg, the pulling clamp, and the main container. The distance from the starting point at the steady edge to the starting point at the pulling edge (L0) is 5 cm. The peg is marked with 0.1% strain intervals. The grooves in the pulling clamp ensures the membrane is intact and stable during the strain

0.1% strain. A complete rotation of the peg is approximately equal to 1% strain. Thus, the change in length or ΔL is 5 mm. Another starting point is marked on top of the container at the pulling end. The pulling camp has multiple grooves to keep the membrane intact while pulling. At 0% strain, the pulling clamp is in line with the starting point marked at the side of the main container, and the 0% mark on the peg is in line with the starting point marked on top of the main container (Fig. 1a, b).

240

Amanda C. Daulagala et al.

In order to validate that the stretching platform was applying a uniform stretch field to our cell cultures, we used standard two-dimensional digital image correlation (DIC) techniques. Briefly, culture membranes were marked by a scattered pattern of pencil shavings and imaged before and after 2.5% and 5% strain (Fig. 2a). Images were then fed into an open-source, MATLABbased DIC algorithm (Ncorr.com) [41], which automatically registers each deformed image to the reference (undeformed) image, identifies particle patterns, measures displacements of particles between deformation steps, and then calculates the full deformation gradient tensor (F) and resulting Engineering strains (defined as the stretch ratio – 1, and corresponding to the macro-scale Tensile strain from Eq. 1 above). We repeated all strain calculations and averaged across three replicates (Fig. 2b, c). 3.2

Cell Culture

1. Wash cut membranes and tweezers with 70% Ethanol and leave in the cell culture hood under UV light on for 20 min until Ethanol is dried out. 2. Switching to bright light, carefully place the membranes inside a 150 mm cell culture dish using tweezers (see Notes 2 and 3). 3. Cover the membrane surfaces with diluted Laminin solution [42] in HBSS as calculated (see Note 1) and leave in the cell culture hood for 1 h (see Notes 4 and 5). 4. Remove the excess fluid and air dry the coated surfaces for 10 min. 5. Evenly plate 10.0  106 Caco2 cells using cell culture medium to homogeneously cover the whole 150 mm plate and leave the plate in the cell culture incubator at 37  C with 5% CO2 until confluent (see Note 6).

3.3 Cell Stretching and Fixing Process

1. Autoclave the glass containers before the experiment begins. 2. Wash the disassembled stretching apparatuses with 70% ethanol and leave in the cell culture hood with UV light on for 20 min until Ethanol is dried out (for different strain conditions, different apparatuses are required). 3. Bring the confluent cell culture dish inside the cell culture hood. 4. Wash the cells with PBS once and add fresh medium. 5. Carefully place one membrane with cells at a time in each of pulling clamp of the apparatus using tweezers. 6. Place the pulling clamp with the membrane inside the main container and drag until the starting point is met by rotating the peg. It is important that the scale of the peg is at zero when the pulling clamp is at the starting point. 7. Lock the free end of the membrane to the steady end using the fasteners.

A Simple Stretch Assay to Study Strain on Epithelial Cells

241

Fig. 2 Experimental platform stretching quantification. (a) Pencil shavings were scattered throughout the membranes and were put under 2.5% and 5% strain with 0% being the resting condition. Engineering strain (¼ stretch ratio – 1) along the axis of the stretching device (x/horizontal direction) was quantified by tracking fiduciary markers before and after 2.5% (b), and 5% (c) strains, measuring the resulting displacements, and calculating strains accordingly. Each strain condition was repeated three times, and, in all cases, the resulting strain fields showed little to no spatial variation across x/horizontal and y/vertical positions. As expected, transverse strains (y/vertical direction) and shear strains were very small relative to axial strains (data not shown)

242

Amanda C. Daulagala et al.

8. Turn the peg until the desired strain is met. For example, a 2.5 turn of the peg approximates to 2.5% strain, and the starting point is 0% strain. 9. Place the apparatuses with membranes inside the glass container (two apparatuses can fit in one container) and fill cell culture media until the membranes are fully covered (approximately 100 mL). 10. Place the closed glass container inside the cell culture cell hood at 37  C with 5% CO2 for 2 h. 11. After 2 h, while the apparatuses are inside the glass container, remove media and wash the membranes with PBS once (see Note 7). 12. Add ice cold methanol into the glass container until the membranes are fully covered and place the glass container at 20  C for 8 min. 13. Remove the methanol and wash membranes three times with PBS. 14. Carefully remove membranes from the apparatuses and place in a new 150 mm dish with PBS (see Note 8). 3.4 Immunofluorescence Staining

1. Block the membranes with blocking reagent for 1 h at room temperature and stain with primary antibodies diluted in antibody diluent overnight at 4  C (for processing of multiple strips see Note 9). 2. Wash three times with PBS and stain with fluorescently labeled secondary antibodies diluted in antibody diluent for 1 h at room temperature. 3. Wash two times with PBS, co-stain with DAPI diluted in PBS, and wash once again with PBS. 4. Add drops of mounting solution on to the mounting slides and place the membranes with cells side up. 5. Add a drop of mounting solution on top of the membrane and place coverslip on top. 6. Seal the edges of coverslip with nail polish and dry overnight at room temperature in the dark.

3.5 Imaging and Analysis

1. Image the slides using confocal microscopy – for this study a Leica SP5 confocal microscope was used with 63 objective and an additional 1.5 zoom. Z-stacks are 0.5 μ in thickness. 2. Open the Z-stacks in ImageJ software [43] and create maximum projections of Z-stacks. 3. Adjust image settings to using Image ! Brightness Contrast option. It should be noted that once the brightness and contrast parameters are set, they should be applied to all the images in an experiment.

A Simple Stretch Assay to Study Strain on Epithelial Cells

243

Table 1 E-cadherin junctional and cytoplasmic immunofluorescence intensity units of Caco2 cells under different conditions of strain (n ¼ 3 representative fields; mean  standard deviation) Strain (%)

Junctional

Cytoplasmic

Junctional/cytoplasmic ratio

0

87.498  12.634

21.935  3.918

3.989  0.715

2.5

31.350  4.910

34.494  7.632

0.934  0.205

0

183.363  5.634

57.979  3.171

3.171  0.245

5

70.472  10.470

65.731  11. 337

1.076  0.029

4. Set the measurement parameters by Analyze ! Set measurements ! Click on Area and Mean gray value. 5. Select the “Free hand line” option and draw a line of a specific length (~10 μm in this experiment) along the membrane. 6. Select Analyze ! Measure. This will provide the area and the mean color value. 7. Repeat steps 5 and 6 for at least additional three times on different areas of the Z-stack image to obtain a representative value. Repeat these steps as necessary to measure cytoplasmic and nuclear color values. It is important to keep the length of the line consistent between the Z-stacks. Table 1, Figs. 3 and 4 show results obtained through the procedure described above. There are significant changes in the E-cadherin localization at 2.5% and 5% strain as measured from the obtained immunofluorescence images (Fig. 3). At both strains E-cadherin junctional localization was decreased, while the cytoplasmic localization was increased (Table 1) resulting in overall decreased junctional/cytoplasmic ratio (Fig. 4).

4

Notes 1. Calculations for Laminin dilution can be performed as follows: area of one strip is 22 cm2 (2 cm  11 cm). Thus, 33 μg (22 cm2  1.5 μg/1 cm2) of Laminin is required to coat one strip. Considering the original concentration of Laminin vial is 1.5 mg/mL and 1 mL of HBSS is required to cover the area of one strip, the dilution is 22 μL of Laminin in 1 mL of HBSS. (33 μg/1.5 mg/mL). 2. Generally, up to three 2 cm  11 cm membranes can be placed inside one 150 mm dish. If more membranes are placed, the area of the membranes that touches the bottom of dish will be decreased. This will negatively affect coating the membranes homogeneously.

244

Amanda C. Daulagala et al.

Fig. 3 E-cadherin localization is affected by strain. Caco2 cells were fixed at different strains and were stained for E-cadherin by immunofluorescence. Each strain was repeated three times with a 0% control. Representative images shown here, where E-cadherin primarily localizes at areas of cell-cell contact at the 0% resting condition. However, at 2.5% (a) and 5% (b) strains, localization of E-cadherin at areas of cell-cell contact is significantly reduced, while the cytoplasmic localization is increased. At 5% strain, a perinuclear E-cadherin localization was also observed, which was included in the cytoplasmic fraction for the calculations

3. Membranes should never dry and must be kept in medium or PBS at all times after fixation. Membranes should only be dried completely under the UV light, right before beginning of cell culture. 4. Sometimes, the diluted Laminin solution may leak out from the membranes into the cell culture dish bottom. This will negatively affect the membrane coating. In this situation, place the membranes in the bottom of the cell culture dish and cover the whole 150 mm plate with diluted Laminin for an even distribution. This will allow for a homogenous coating of the membranes.

A Simple Stretch Assay to Study Strain on Epithelial Cells

245

Fig. 4 The ratio of junctional vs cytoplasmic E-cadherin localization is decreased upon application of strain in Caco2 cells. Graphical representation of the E-cadherin junctional and cytoplasmic immunofluorescence quantification of Caco2 cells under different conditions of strain presented in Table 1. Upon application of strain, the ratio of junctional to cytoplasmic localization of E-cadherin is significantly reduced (n ¼ 3 representative fields; mean  standard deviation; *P < 0.05, student’s t-test)

5. We have identified Laminin as the substrate where the junctional localization of E-cadherin agrees with published work in well-differentiated Caco2 cells and tissues by us and others [29, 42, 44–46]. Nevertheless, other ECM proteins such as Collagens and Fibronectin can be used according to different cell types. 6. 10  106 [47] is half the cells at confluency of a 150 mm dish according to Fisher Scientific protocols. If cells are not attaching and growing properly, the seeding density can be changed accordingly. It is also important to note that seeding density varies depending on the cell line. 7. Do not accidentally stretch the membranes while handling and placing inside and taking out of the apparatus. 8. Fixed membranes can be immersed in PBS, and stored at 4  C until ready to be stained. 9. For multiple stains, membranes can be cut in 3–4 pieces accordingly, while they are still in PBS, before the blocking step.

Acknowledgments This work was supported by: NIH P20 GM130457-01A1 (to AK); NIH P20 GM103444 (SC BioCRAFT pilot Award to AK); Concern Foundation’s Conquer Cancer Now Award (to AK); NIH P20 GM121342 (to WJR).

246

Amanda C. Daulagala et al.

References 1. Miller CJ, Davidson LA (2013) The interplay between cell signalling and mechanics in developmental processes. Nat Rev Genet 14 (10):733–744 2. Barnes LA, Marshall CD, Leavitt T, Hu MS, Moore AL, Gonzalez JG, Longaker MT, Gurtner GC (2018) Mechanical forces in cutaneous wound healing: emerging therapies to minimize scar formation. Adv Wound Care (New Rochelle) 7(2):47–56 3. Broders-Bondon F, Nguyen Ho-Bouldoires TH, Fernandez-Sanchez M-E, Farge E (2018) Mechanotransduction in tumor progression: the dark side of the force. J Cell Biol 217(5):1571–1587 4. Kalli M, Stylianopoulos T (2018) Defining the role of solid stress and matrix stiffness in cancer cell proliferation and metastasis. Front Oncol (2234-943X (Print)) 5. Kim ER (2014) Colorectal cancer in inflammatory bowel disease: the risk, pathogenesis, prevention and diagnosis. World J Gastroenterol 20(29):9872 6. Gordon IO, Agrawal N, Willis E, Goldblum JR, Lopez R, Allende D, Liu X, Patil DY, Yerian L, El-Khider F, Fiocchi C, Rieder F (2018) Fibrosis in ulcerative colitis is directly linked to severity and chronicity of mucosal inflammation. Aliment Pharmacol Ther 47 (7):922–939 7. Basson MD, Li GD, Hong F, Han O, Sumpio BE (1996) Amplitude-dependent modulation of brush border enzymes and proliferation by cyclic strain in human intestinal Caco-2monolayers. J Cell Physiol 168(2):476–488 8. Ali MY, Saif MTA (2014) Substrate stiffness mediated metastasis like phenotype of colon cancer cells is independent of cell to gel adhesion. Cell Mol Bioeng 7(4):532–543 9. Ferna´ndez-Sa´nchez ME, Barbier S, Whitehead J, Be´alle G, Michel A, LatorreOssa H, Rey C, Fouassier L, Claperon A, Brulle´ L, Girard E, Servant N, Rio-Frio T, Marie H, Lesieur S, Housset C, Gennisson J-L, Tanter M, Me´nager C, Fre S, Robine S, Farge E (2015) Mechanical induction of the tumorigenic β-catenin pathway by tumour growth pressure. Nature 523(7558):92–95 10. Ro¨per J-C, Mitrossilis D, Stirnemann G, Waharte F, Brito I, Fernandez-Sanchez M-E, Baaden M, Salamero J, Farge E (2018) The major β-catenin/E-cadherin junctional binding site is a primary molecular mechanotransductor of differentiation in vivo. Elife 7 11. Ciasca G, Papi M, Minelli E, Palmieri V, De Spirito M (2016) Changes in cellular mechanical properties during onset or progression of

colorectal cancer. World J Gastroenterol 22 (32):7203–7214 12. Thompson WR, Scott A, Loghmani MT, Ward SR, Warden SJ (2016) Understanding mechanobiology: physical therapists as a force in mechanotherapy and musculoskeletal regenerative rehabilitation. Phys Ther 96(4):560–569 13. Rawlinson SCF (2017) Mechanical responsiveness of distinct skeletal elements: possible exploitation of low weight-bearing bone. In: Rawlinson SCF (ed) Mechanobiology: exploitation for medical benefit. Wiley, Hoboken, pp 131–138 14. Kalli M, Stylianopoulos T (2018) Defining the role of solid stress and matrix stiffness in cancer cell proliferation and metastasis. Front Oncol 8:55 15. Atcha H, Davis CT, Sullivan NR, Smith TD, Anis S, Dahbour WZ, Robinson ZR, Grosberg A, Liu WF (2018) A low-cost mechanical stretching device for uniaxial strain of cells: a platform for pedagogy in mechanobiology. J Biomech Eng 140(8) 16. Shao Y, Tan X, Novitski R, Muqaddam M, List P, Williamson L, Fu J, Liu AP (2013) Uniaxial cell stretching device for live-cell imaging of mechanosensitive cellular functions. Rev Sci Instrum 84(11):114304 17. Seriani S, Del Favero G, Mahaffey J, Marko D, Gallina P, Long CS, Mestroni L, Sbaizero O (2016) The cell-stretcher: a novel device for the mechanical stimulation of cell populations. Rev Sci Instrum 87(8):084301 18. USA, S. Cell Stretching Systems (2020) https://strexcell.com/cell-stretching-system/ 19. Hoffman BD, Grashoff C, Schwartz MA (2011) Dynamic molecular processes mediate cellular mechanotransduction. Nature 475 (7356):316–323 20. Lecuit T, Yap AS (2015) E-cadherin junctions as active mechanical integrators in tissue dynamics. Nat Cell Biol 17(5):533–539 21. Lecuit T (2010) Alpha-catenin mechanosensing for adherens junctions. Nat Cell Biol 12 (6):522–524 22. Schwartz MA (2010) Integrins and extracellular matrix in mechanotransduction. Cold Spring Harb Perspect Biol 2(12):a005066 23. Ross TD, Coon BG, Yun S, Baeyens N, Tanaka K, Ouyang M, Schwartz MA (2013) Integrins in mechanotransduction. Curr Opin Cell Biol 25(5):613–618 24. Borghi N, Sorokina M, Shcherbakova OG, Weis WI, Pruitt BL, Nelson WJ, Dunn AR (2012) E-cadherin is under constitutive actomyosin-generated tension that is increased

A Simple Stretch Assay to Study Strain on Epithelial Cells at cell-cell contacts upon externally applied stretch. Proc Natl Acad Sci U S A 109 (31):12568–12573 25. Benham-Pyle BW, Pruitt BL, Nelson WJ (2015) Cell adhesion. Mechanical strain induces E-cadherin-dependent Yap1 and betacatenin activation to drive cell cycle entry. Science 348(6238):1024–1027 26. Daulagala AC, Bridges MC, Kourtidis A (2019) E-cadherin beyond structure: a signaling hub in Colon homeostasis and disease. Int J Mol Sci 20(11) 27. Kourtidis A, Lu R, Pence LJ, Anastasiadis PZ (2017) A central role for cadherin signaling in cancer. Exp Cell Res 358(1):78–85 28. Kourtidis A, Necela B, Lin WH, Lu R, Feathers RW, Asmann YW, Thompson EA, Anastasiadis PZ (2017) Cadherin complexes recruit mRNAs and RISC to regulate epithelial cell signaling. J Cell Biol 216(10):3073–3085 29. Kourtidis A, Ngok SP, Pulimeno P, Feathers RW, Carpio LR, Baker TR, Carr JM, Yan IK, Borges S, Perez EA, Storz P, Copland JA, Patel T, Thompson EA, Citi S, Anastasiadis PZ (2015) Distinct E-cadherin-based complexes regulate cell behaviour through miRNA processing or Src and p120 catenin activity. Nat Cell Biol 17(9):1145–1157 30. Nair-Menon J, Daulagala AC, Connor DM, Rutledge L, Penix T, Bridges MC, Wellslager B, Spyropoulos DD, Timmers CD, Broome AM, Kourtidis A (2020) Predominant distribution of the RNAi machinery at apical Adherens junctions in colonic epithelia is disrupted in Cancer. Int J Mol Sci 21(7) 31. Chen CH, Marymont JV, Huang MH, Geyer M, Luo ZP, Liu X (2007) Mechanical strain promotes fibroblast gene expression in presence of corticosteroid. Connect Tissue Res 48(2):65–69 32. Manuyakorn W, Smart DE, Noto A, Bucchieri F, Haitchi HM, Holgate ST, Howarth PH, Davies DE (2016) Mechanical strain causes adaptive change in bronchial fibroblasts enhancing Profibrotic and inflammatory responses. PLoS One 11(4):e0153926 33. Matsumoto T, Yung YC, Fischbach C, Kong HJ, Nakaoka R, Mooney DJ (2007) Mechanical strain regulates endothelial cell patterning in vitro. Tissue Eng 13(1):207–217 34. Zeiger AS, Liu FD, Durham JT, Jagielska A, Mahmoodian R, Van Vliet KJ, Herman IM (2016) Static mechanical strain induces capillary endothelial cell cycle re-entry and sprouting. Phys Biol 13(4):046006 35. Lea T (2015) Caco-2 cell line. In: Verhoeckx K, Cotter P, Lo´pez-Expo´sito I, Kleiveland C, Lea T, Mackie A, Requena T, Swiatecka D,

247

Wichers H (eds) The impact of food bioactives on health. Springer, Cham, pp 103–111 36. Grasset E, Bernabeu J, Pinto M (1985) Epithelial properties of human colonic carcinoma cell line Caco-2: effect of secretagogues. Am J Physiol 248(5 Pt 1):C410–C418 37. Hidalgo IJ, Raub TJ, Borchardt RT (1989) Characterization of the human colon carcinoma cell line (Caco-2) as a model system for intestinal epithelial permeability. Gastroenterology 96(3):736–749 38. Samak G, Gangwar R, Crosby LM, Desai LP, Wilhelm K, Waters CM, Rao R (2014) Cyclic stretch disrupts apical junctional complexes in Caco-2 cell monolayers by a JNK-2-, c-Src-, and MLCK-dependent mechanism. Am J Physiol Gastrointest Liver Physiol 306(11): G947–G958 39. Formlabs (2020) https://formlabs.com/ software/ 40. Ling SJ, Sanny J, Moebs W (eds) University Physics.Open Stax. Rice University, Houston 41. Blaber J, Adair B, Antoniou A (2015) Ncorr: open-source 2D digital image correlation Matlab software. Exp Mech 55:1105–1122 42. Basson MD, Turowski G, Emenaker NJ (1996) Regulation of human (Caco-2) intestinal epithelial cell differentiation by extracellular matrix proteins. Exp Cell Res 225(2):301–305 43. Schindelin J, Arganda-Carreras I, Frise E, Kaynig V, Longair M, Pietzsch T, Preibisch S, Rueden C, Saalfeld S, Schmid B, Tinevez JY, White DJ, Hartenstein V, Eliceiri K, Tomancak P, Cardona A (2012) Fiji: an opensource platform for biological-image analysis. Nat Methods 9(7):676–682 44. Kourtidis A, Anastasiadis PZ (2016) PLE KHA7 defines an apical junctional complex with cytoskeletal associations and miRNAmediated growth implications. Cell Cycle 15 (4):498–505 45. Jokhadar SˇZ, Sˇusˇtar V, Svetina S, Batista U (2009) Time lapse monitoring of CaCo-2 cell shapes and shape dependence of the distribution of integrin β1 and F-actin on their basal membrane. Cell Commun Adhes 16 (1–3):1–13 46. Zhang X, Cromwell JW, Kunjummen BD, Yee D, Garcia-Aguilar J (2003) The α2 and α3 integrins are required for morphologic differentiation of an intestinal epithelial cell line. Surgery 133(4):429–437 47. ThermoScientific (2020) Useful numbers for cell culture. https://www.thermofisher.com/ us/en/home/references/gibco-cell-culturebasics/cell-culture-protocols/cell-culture-use ful-numbers.html

Methods in Molecular Biology (2021) 2367: 249–271 DOI 10.1007/7651_2021_389 © Springer Science+Business Media, LLC 2021 Published online: 09 April 2021

Differentiating Between Tight Junction-Dependent and Tight Junction-Independent Intestinal Barrier Loss In Vivo Sandra D. Chanez-Paredes, Shabnam Abtahi, Wei-Ting Kuo, and Jerrold R. Turner Abstract The intestinal barrier is an essential component of innate host defense. The single layer of epithelial cells that line the intestine must balance barrier function with both active, transcellular and diffusive, paracellular transport. Tight junctions, which link adjacent cells, form a selectively permeable seal that defines both paracellular transport and barrier properties. Molecules can cross tight junctions by either of two distinct routes, termed pore and the leak pathways, that differ in capacity, charge-selectivity, size-selectivity, and responses to physiological and pathophysiological stimuli. A third intestinal permeability route, the unrestricted pathway, reflects loss of the epithelial barrier, as occurs with mucosal damage, is independent of paracellular and transcellular pathways, and is neither charge- nor size-selective. The most commonly used approach for measuring intestinal permeability in vivo involves gavage of FITC-4 kDa dextran and analysis of the quantity recovered in serum. Unfortunately, this method cannot distinguish between leak and unrestricted pathways, as 4 kDa dextran can cross both. Moreover, 4 kDa dextran is too large to cross the pore pathway and, therefore, provides no information regarding this paracellular flux route. Here we describe a multiplex method that allows simultaneous, independent analysis of each pathway. Key words Barrier dysfunction, Claudins, Gavage, Intestinal permeability, Ion flux, Leak pathway, Macromolecular flux, Paracellular flux, Pore pathway, Tight junction, Unrestricted pathway

1

Introduction The intestinal epithelial barrier is created by the single layer of epithelial cells that line the gastrointestinal tract [1]. This interface, which is the largest mucosal surface within the body, is the primary site at which the developing immune system interacts with foreign elements, including the microbiome [2–5]. Epithelial cells, whose lipid membranes are impermeant to many types of foreign materials, modulate contact between luminal materials and mucosal immune cells by regulating flux across the paracellular, or shunt, pathway [6–10] and by transcellular transport [11, 12]. Although barrier function is emphasized most frequently, tight junctions within many organs, including the gut, must also be

249

250

Sandra D. Chanez-Paredes et al.

Fig. 1 Mechanisms of intestinal permeability. Paracellular flux across tight junctions occurs by two different routes termed pore and the leak pathways. The pore pathway is a high-capacity, charge-selective route that regulates the passage of small molecules less than
8 Å in diameter. The leak pathway is a low-capacity, non-charge selective route traversed by molecules up to
100 Å in diameter. When epithelial damage occurs the tight junction-independent, size-nonselective, charge-nonselective unrestricted pathway becomes the dominant route of intestinal permeability

selectively permeable. Paracellular ion and water flux are mediated by small trans-tight junction channels created by some members of the claudin protein family [13–16]. The critical nature of this selective permeability is demonstrated by perinatal death of knockout mice lacking claudins 2 and 15, which are essential for intestinal paracellular Na+ flux [17]. This high-capacity flux route (Fig. 1) has been termed the pore pathway and is both charge- and sizeselective [1, 18]. The charge-selectivity of channels formed by different claudins varies, but, most have maximal diameters of 6–8 Å [19–24]. Functional modulation of the intestinal pore pathway occurs during development and disease as a consequence of regulated claudin isoform expression. For example, claudin-2 is normally expressed at high levels prior to weaning but is replaced by claudin-15 after transition to a solid diet [25, 26]. Claudin-2 expression is, however, elevated once again in the context of intestinal disease, including inflammatory and infectious disorders

Multiplex Intestinal Permeability Analysis

251

[27–30]. Although incompletely studied, this claudin-2 upregulation can be selectively induced by IL-13 and IL-22 [27, 28, 31] and can both promotes pathogen clearance, which is beneficial during enteric infection [28], and amplify immune responses, which is detrimental in immune-mediated colitis [32]. In contrast to the pore pathway, the leak pathway (Fig. 1) is not charge-selective and has a relatively low capacity. Although it is sizeselective, the maximum diameter of the leak pathway is estimated to be ~100 Å, far greater than the pore pathway. Thus, small proteins and bacterial products, e.g., albumin and lipopolysaccharide, can traverse the epithelial barrier via the leak pathway but intact bacteria and viruses cannot. Intestinal epithelial leak pathway permeability is primarily regulated via the cytoskeleton as a result of myosin light chain kinase activation, which leads to phosphorylation of perijunctional myosin II regulatory light chain [33–36]. Unlike the pore pathway, a physical channel that defines the leak pathway has not been identified. The final flux route that must be considered is the tight junction-independent unrestricted pathway (Fig. 1). This route exists only in the context of epithelial damage, e.g., when there are no intercellular junctions due to epithelial cell loss. As might be expected, the unrestricted pathway has a high-capacity and is neither size- nor charge-selective [37]. In vivo assessment of intestinal permeability in rodents has relied on oral gavage of tracer molecules and subsequent detection of these within the blood [38–44]. These studies have primarily employed low molecular weight, i.e., 3 kDa or 4 kDa dextrans, conjugated to fluorescent molecules such as fluorescein, and fluorescein itself has been used as a smaller probe [45]. Because fluorescein and small dextrans cross both leak and unrestricted pathways, increased serum recovery cannot distinguish between enhanced leak pathway permeability and mucosal damage, i.e., unrestricted pathway flux. Moreover, because these probes are too large to cross the pore pathway, they provide no information on permeability of this paracellular route. Thus, although many studies have noted a correlation between claudin-2 upregulation and increased flux of FITC-4 kDa dextran and therefore concluded that claudin-2 upregulation mediates 4 kDa dextran flux [46–49], this is incorrect. This protocol uses 3 separate probes: creatinine (6 Å diameter), FITC-4 kDa dextran (28 Å diameter), and rhodamine-70 kDa dextran (120 Å diameter). Creatinine can traverse all three pathways, 4 kDa dextran is restricted to leak and unrestricted pathways, and 70 kDa dextran is limited to the unrestricted pathway. Thus, an increase in 4 kDa dextran flux accompanied by comparable increases in 70 kDa dextran flux can be recognized as a consequence of enhanced flux across the unrestricted pathway. In this case, where both probes can pass through the unrestricted pathway,

252

Sandra D. Chanez-Paredes et al.

it is important to recognize that increases in 4 kDa dextran serum recovery may be greater than those for 70 kDa dextran due to sieving properties, i.e., a very small object passes through a large channel more readily than an object only slightly smaller than the channel. In contrast, increased 4 kDa dextran flux without changes in 70 kDa dextran flux reflects leak pathway permeability. Altered creatinine flux must similarly be interpreted in the context of 4 kDa and 70 kDa dextran recoveries. By using these 3 probes, it is possible to discriminate between tight junction-dependent pore and leak pathways as well as the tight junction-independent unrestricted pathway as mediators of altered intestinal permeability in vivo.

2 2.1

Materials Reagents

1. Anhydrous creatinine; Millipore-Sigma C4255. 2. Fluorescein isothiocyanate (FITC)-4 kDa dextran; MilliporeSigma FD4. 3. Rhodamine B isothiocyanate-70 kDa dextran; Millipore-Sigma R9379. 4. Enzyme-based creatinine assay kit (see Note 1). 5. 10 kDa spin columns (see Note 2). 6. Ultrapure (18 MΩ) water. 7. Sterile PBS: 140 mM NaCl, 10 mM sodium phosphate, pH 7.4. 8. 100 22 Ga stainless-steel gavage needle with 1.25 mm ball tip; Braintree Scientific N-PK 002. 9. Sterile 0.22-μm syringe filter. 10. Sterile 10 ml and 1 ml syringes. 11. Sterile 50 ml tubes. 12. Sterile disposable 1-000-0500.

glass

micropipettes;

Drummond

13. Micro plasma collection tubes with 3.2% citrate anticoagulant; Sarstedt, 41.1506.002 (see Note 3). 14. 96-well solid black polystyrene microplates (see Note 4). 15. Sterile anesthetic solution: 10 mg/ml ketamine, 1 mg/ml xylazine in 0.9% saline. 2.2

Equipment

1. Microcentrifuge. 2. Fluorescent plate reader (see Note 5). 3. 37  C water bath.

Multiplex Intestinal Permeability Analysis

3

253

Methods

3.1 Probe Preparation

1. Calculate the total volume required and the quantity needed for each probe. The probe stock, prepared in water, contains 100 mg/ml creatinine, 80 mg/ml FITC-4 kDa dextran, and 40 mg/ml rhodamine-70 kDa dextran (see Note 6). 2. Add probes and ultrapure water to a 50 ml conical tube and place in 37  C water bath (see Note 7). 3. Incubate the tube in the water bath. Swirl gently every 2–3 min to mix until the probes are completely dissolved, typically ~15 min. 4. Centrifuge the 50 ml tube at 500  g for 5 min. Use a Pasteur pipette to resuspend and solubilize, if possible, any particulate pellet. 5. Filter the probe cocktail into a fresh, sterile 50 ml tube using the 10 ml syringe and 0.22 μm filter. 6. Centrifuge the 50 ml tube at 500  g for 1 min to bring all of the probe solution to the tube bottom. 7. Remove 20 μl of probe for later use in standard preparation. 8. Gavage mice within 2–3 h of preparation. Do not store for later use.

3.2 Gavage (See Notes 8 and 9)

1. Select mice for study (see Note 10). 2. In order to clear gastrointestinal contents, place mice in a fresh cage, without chow or bedding (see Note 11), 3 h prior to gavage. Water should be provided during this time. 3. Draw 0.25 ml of gavage solution into a 1 ml syringe and attach a clean gavage needle. Point the needle upward and gently press the plunger to expel any air. 4. Scruff a mouse by holding the tail with one hand while grasping the skin at the nape of the neck (see Note 12). 5. Gently retract the neck and hold the mouse vertically. 6. Hold the syringe, with needle pointed upward, next to the mouse to assess the distance from the mouth to the xiphoid process (Fig. 2a). This is the distance to which the needle should be inserted into the mouse (see Note 13). 7. Carefully tilt the head of the mouse backward so that the head, body, and esophagus are aligned (see Note 14). 8. Gently push the needle over the tongue and to the back of the oral cavity. The mouse may try to lick the needle; continue advancing the needle until the ball tip touches the pharynx (see Note 15).

254

Sandra D. Chanez-Paredes et al.

Fig. 2 Probe gavage. (a) After restraining the mouse, hold the syringe with the needle pointed upward in order to assess the length to be inserted. (b) The needle should be inserted into the proximal stomach

9. The mouse will start swallowing reflexively. It is important to allow this and refrain from manually forcing the needle beyond the pharynx, which can cause tissue damage (see Note 16). The gavage needle should easily slide along the esophagus (Fig. 2b). 10. Instill probe solution (see Note 17). 11. Record the exact time of gavage (see Note 18). 12. Gently withdraw the gavage needle. 13. Return the mouse to the cage. Water, but not chow, should be provided. 14. Watch the mouse until normal respiration and activity have returned (see Note 19). 15. Repeat with the next mouse (see Note 20). 16. Chow can be returned to the cage 90 min after gavage. 3.3 Blood Collection and Plasma Separation

1. 30 min before blood collection, label plasma collection tubes with the identifier for each mouse and mark the level to which blood should be collected (see Note 21). 2. Anesthetize the mouse (see Note 22) about 2 h and 50 min after gavage (see Note 23). 3. Hold the mouse tightly by the back of the neck, with sufficient force to slightly retract the skin of the head, with the thumb and either the index or middle finger.

Multiplex Intestinal Permeability Analysis

255

Fig. 3 Blood collection. (a) The tail vein and retro-orbital sinus can both be used as sites for blood collection. To retrieve blood from the retro-orbital sinus, a sterile micropipette is placed at the medial canthus and inserted. (b) The volume of blood needed should be marked in advance on the side of each sodium citrate collection tube (1). After blood collection, the tube should be gently inverted to mix the blood and anticoagulant (2). Centrifugation leaves transparent, straw-colored plasma above a cellular pellet (3). If the plasma is red-tinged, hemolysis has occurred and the sample may yield inaccurate results

4. Retract the eyelid and proptose the eye. 5. Place a glass micropipette at the lateral canthus of the eye under the nictitating membrane (Fig. 3a). 6. Gently, press the micropipette against the orbital bone until blood begins to flow. 7. Fill the collection tube to the indicated line (Fig. 3b; see Note 24). 8. Apply slight pressure on the eye after using a clean gauze pad (see Note 25). 9. To prevent coagulation, immediately mix the blood with the buffered sodium citrate solution by inverting the tube 5–10 times. Do not mix vigorously, as this can result in hemolysis (see Note 26). 10. After blood has been collected from all mice, spin the collection tubes at 1500  g for 10 min at room temperature. 11. Transfer the plasma to new tubes. Take care not to disturb the cell pellet. 12. It is best to proceed with analysis immediately (see Note 27). 3.4 Measuring Plasma Fluorescein and Rhodamine B Fluorescence

1. Prepare a standard curve from the probe stock solution (see Note 28). Add 435 μl of ultrapure water to 15 μl of probe stock to create a 1:30 dilution. Add 450 μl of water to 50 μl of the 1:30 dilution to create a 1:300 dilution. Use this to create a 1:3,000 dilution and repeat until 1:30,000, 1:300,000, and 1:3,000,000 dilutions have been prepared. Add 133 μl of water to 67 μl of the 1:30 dilution to create a 1:100 dilution. Add 450 μl of water to 50 μl of the 1:100 dilution to create a 1:1,000 dilution. Dilute this serially to 1:10,000, 1:100,000, and 1:1,000,000.

256

Sandra D. Chanez-Paredes et al.

2. Pipette exactly 100 μl of each standard dilution into one well within the solid black microplate. Include water to determine background for the standard samples (see Note 29). 3. Pipette exactly 100 μl of each plasma sample into one well within the solid black microplate (see Note 30). Include plasma from the mouse, or mice, gavaged with water to determine background for the serum samples. 4. Read fluorescein and rhodamine B fluorescence at Ex/Em of 495 nm/525 nm and 555 nm/585 nm, respectively, at gains of 50, 60, 70, 80, 90 and 100 for fluorescein, and 80, 90, 100, 110, 120 and 130 for rhodamine B (see Note 31). 3.5 Measuring Plasma Creatinine Concentration

1. Prepare a 1 nmol/μl creatinine standard (see Note 32) by diluting 5 μl of the 100 mM creatinine standard provided in the kit with 495 μl of assay buffer (see Notes 33 and 34). 2. Prepare a 0.1 nmol/μl dilution by adding 100 μl of the 1 nmol/μl standard to 900 μl of assay buffer. 3. Use the 0.1 nmol/μl standard to prepare a standard curve. Pipette 150 μl, 144 μl, 138 μl, 132 μl, 126 μl, and 120 μl of assay buffer into successive tubes or microplate wells and add 0 μl, 6 μl, 12 μl, 18 μl, 24 μl, and 30 μl of the 0.1 nmol/μl standard to the wells, respectively, to generate standards containing 0 nmol, 0.2 nmol, 0.4 nmol, 0.6 nmol, 0.8 nmol, and 1.0 nmol per 50 μl. Pipette 50 μl of each standard into successive wells of a solid black microplate (see Note 35). 4. Remove 75 μl of plasma from each well of the microplate and dilute 1:4 with 225 μl of water. 5. Load the diluted plasma into the 10 kDa spin columns and centrifuge at 10,000  g for 10 min at 4  C (see Note 36). 6. Recover 10 μl of filtered plasma and dilute 1:20 by adding 190 μl of room temperature assay buffer. 7. Pipette 50 μl of each 1:20 diluted plasma into a well for the creatinine assay (see Note 37). 8. Prepare the creatinine reaction master mix. For each sample, add 43.6 μl of assay buffer, 2 μl of creatininase, 2 μl of creatinase, 2 μl of enzyme mix, and 0.4 μl of creatinine probe (see Note 38). 9. Add 50 μl of reaction master mix to each sample and standard well (see Note 39). 10. Mix gently using an orbital rotator. 11. Incubate for 1 h at 37  C (see Note 40). 12. Analyze the samples using the plate reader set to Ex/Em of 538 nm/587 nm (see Note 41) with gains of 50, 60, 70, 80, 90, and 100.

Multiplex Intestinal Permeability Analysis

3.6

Calculations

257

1. Select readings to be used for each probe. Use results from the gain setting that result in RFU values of ~200 for the samples from the water-gavaged control mice (see Note 42). 2. Subtract the blank (water) value, as background, from all standards. 3. For each probe, use the background-subtracted standard values and linear regression analysis to generate a straight line defining the relationship between fluorescent signal and probe concentration (see Note 43). 4. Use the regression equation to determine the concentration of each probe in each sample. 5. Subtract the values calculated for the control plasma sample (from the mouse gavaged with water only) from each experimental sample. If more than one control mouse was gavaged with water, the mean of these values should be used (see Note 44). 6. Calculate the mean and standard deviation of each probe’s concentration for the mice in the healthy control, probegavaged group (see Note 45). 7. Normalize the value of each probe in each mouse to the mean of the healthy control group. 8. Use the normalized values to calculate, for each mouse, the ratios of creatinine:70 kDa dextran, 4 kDa dextran:70 kDa dextran, and creatinine:4 kDa dextran (see Note 46). 9. Plot the normalized values for each probe and the three ratios for each mouse.

3.7 Data Interpretation

Ideally, flux data should be assessed quantitatively and the apparent permeability coefficient, Papp, of each probe should be plotted as a function of probe size [50]. This calculation involves, at minimum, 3 variables; the measured probe transport rate, the initial concentration of probe at the apical cell surface, and the surface area assessed. Although transport, i.e., flux, rate can be measured, neither the initial concentration at the apical surface nor the surface area can be determined in this assay. Thus, in contrast to Ussing chamber analyses, Papp cannot be accurately calculated and plotted as a function of probe size [50–53]. We, therefore, typically present data normalized to values obtained in healthy control mice. It is, nevertheless, important to consider flux of each probe in concert with fluxes of the other probes. After normalization, both individual values for each probe as well as ratios should be assessed. For example, increased creatinine flux without changes in 4 kDa or 70 kDa dextran fluxes would result in increased ratios of creatinine-to-70 kDa dextran and creatinine-to-4 kDa dextran but should have no effect on the 4 kDa dextran-to-70 kDa dextran ratio. These changes can be seen early in

258

Sandra D. Chanez-Paredes et al.

Fig. 4 Pore pathway upregulation after C. rodentium infection. (a) Colonic histology before and 2 days after C. rodentium infection. Bar ¼ 20 μm. (b) Pseudocolor image showing that C. rodentium infection increases both the number of cells expressing claudin-2 and the amount of claudin-2 expressed by each cell (arrows).

Multiplex Intestinal Permeability Analysis

259

the course of C. rodentium infection (Fig. 4) [28]. This is accompanied by selective increases in expression of claudin-2, which forms paracellular channels that mediate pore pathway flux [1, 13–16, 28]. Notably, perijunctional myosin II regulatory light chain phosphorylation is not affected at this early time point (Fig. 4). In contrast to C. rodentium infection, flux across the leak pathway is dramatically upregulated in response to acute T cell activation (Fig. 5). This TNF-dependent permeability change is accompanied by increases in both creatinine and 4 kDa dextran flux without any change in 70 kDa dextran flux. The creatinine-to70 kDa dextran and 4 kDa dextran-to-70 kDa dextran ratios are both increased, but, at most, only modest changes are seen in the creatinine-to-4 kDa dextran ratio. In this example, pore pathway permeability may be upregulated but cannot be readily detected, as creatinine and 4 kDa dextran can both traverse the leak pathway, and increased creatinine flux is therefore expected in isolated leak pathway upregulation. Thus, even if claudin-2 expression and pore pathway permeability were increased, those changes would be functionally overwhelmed by the magnitude of leak pathway permeability increases. Small increases in the creatinine-to-4 kDa dextran ratio are expected due to the relative sizes of these probes and leak pathway sieving properties, but marked increases could indicate functionally significant increases in permeability of both pore and leak pathways. In vivo, TNF-induced diarrhea, which peaks at 2–3 h and resolves within 5 h [33, 34], is accompanied by increases in myosin II regulatory light chain phosphorylation (Fig. 5). Claudin-2 expression is not affected, consistent with other data indicating that claudin-2 upregulation does not occur until 12–18 h after administration of exogenous cytokines [28, 31, 32]. Finally, the severe mucosal ulceration induced by dextran sulfate sodium (DSS) increases flux of all 3 probes (Fig. 6) but only small changes in the calculated ratios. Because the small probes creatinine and 4 kDa dextran, cross the unrestricted pathway more readily than 70 kDa dextran, changes in creatinine-to70 kDa dextran and 4 kDa dextran-to-70 kDa dextran fluxes can ä Fig. 4 (continued) Bar ¼ 20 μm. (c) Merged image of the same fields as B showing claudin-2 (red), phosphorylated MLC (white), Na-K ATPase (green), and nuclei (blue). Bar ¼ 20 μm. (d) In the absence of infection, flux of all three probes is limited. (e) C. rodentium infection increases pore pathway permeability, which can be measured as creatinine flux. (f) Creatinine flux before and 2 days after C. rodentium infection. (g) 4 kDa dextran flux before and 2 days after C. rodentium infection. H. 70 kDa dextran flux before and 2 days after C. rodentium infection. (i) The creatinine-to-70 kDa dextran ratio is increased, indicating pore pathway upregulation in this context. (j) The 4 kDa dextran-to-70 kDa dextran ratio is unchanged. (k) The creatinine-to4 kDa dextran ratio is increased, indicating that the increase in creatinine flux does not reflect leak pathway upregulation. Mean  SD, *p < 0.05

260

Sandra D. Chanez-Paredes et al.

Fig. 5 Leak pathway upregulation after T cell activation. (a) Jejunal histology before and 3 h after anti-CD3 treatment. Bar ¼ 20 μm. (b) Pseudocolor image showing increased phosphorylated MLC (phosphoMLC) within epithelial cells, particularly at the perijunctional actomyosin ring (arrows). MLC phosphorylation is also

Multiplex Intestinal Permeability Analysis

261

be seen, but are smaller than increases in creatinine or 4 kDa dextran fluxes (Fig. 6). This pattern of change reflects increased unrestricted pathway permeability. Although simultaneous increases in pore and leak pathway permeabilities may occur at sites without epithelial damage, these cannot typically be measured, as their impact is masked by the large changes in unrestricted pathway permeability and, as a result of sieving properties, much greater increases in flux of creatinine and 4 kDa dextran across the unrestricted pathway. Thus, when evaluating studies in which only 4 kDa dextran was used as a probe, large recovery increases should be interpreted as most likely representing mucosal damage rather than tight junction regulation. A final consideration is that these measurements can be affected by intestinal and extra-intestinal factors, including intestinal tissue architecture [55, 56], luminal flow rate [51, 57], and probe clearance by the kidneys and mononuclear phagocyte system. Changes in any of these factors may, therefore, impact measurements independent of changes in tight junction permeability or mucosal damage. Thus, while in vivo multiplex analysis of intestinal permeability is a powerful tool, it is critical that data be interpreted in the context of the underlying biology.

4

Notes 1. The colorimetric alkaline picrate-creatinine (Jaffe) reaction [58] can also be used to measure creatinine. However, this reaction is not entirely specific, and several substances present in blood, including glucose, ketones, protein, and bilirubin, can affect results [59]. We have, therefore, used an enzymatic assay in which creatininase (EC 3.5.2.10) converts creatinine to creatine, and creatinase (EC 3.5.3.3) generates sarcosine. Sarcosine is then is oxidized by sarcosine oxidase (EC 1.5.3.1) to produce hydrogen peroxide, glycine, and 5,10-CH2-tetrahydrofolate. The hydrogen peroxide then interacts with a colorimetric/fluorometric probe. Commercial versions of this kit are

ä Fig. 5 (continued) increased within vascular endothelium (asterisks). Bar ¼ 20 μm. (c) Merged image of the same fields as B showing claudin-2 (red), phosphorylated MLC (white), Na-K ATPase (green), and nuclei (blue). Bar ¼ 20 μm. (d) Before T cell activation, flux of all three probes is limited. (e) T cell activation increases leak pathway permeability, which can be measured as 4 kDa dextran flux. (f) Creatinine flux before and 3 h after T cell activation. (g) 4 kDa dextran flux before and 3 h after T cell activation. (h) 70 kDa dextran flux before and 3 h after T cell activation. (i) The creatinine-to-70 kDa dextran ratio is increased. (j) The 4 kDa dextran-to70 kDa dextran ratio is increased, indicating leak pathway upregulation. (k) The creatinine-to-4 kDa dextran ratio is not changed, indicating that the increase in creatinine flux reflects leak pathway upregulation. Mean  SD, **p < 0.01, *p < 0.05

262

Sandra D. Chanez-Paredes et al.

Fig. 6 Intestinal unrestricted pathway permeability increases after DSS-induced damage. (a) Colonic histology before and after 7 days of DSS treatment. Note the regenerative crypt epithelium (arrow) and area of ulceration (asterisk) associated with DSS-induced damage. Bar ¼ 20 μm. (b) Merged image showing CD3-positive (red) T cell infiltration after DSS-induced damage (asterisk) and early re-epithelialization of the ulcer (arrow). ZO-1

Multiplex Intestinal Permeability Analysis

263

available from Millipore-Sigma (MAK079), Abcam (ab65340), and other vendors. 2. High protein concentrations can interfere with the creatinine assay. Plasma should be deproteinized using 10 kDa spin columns, e.g., Millipore-Sigma CLS431478 or Abcam ab93349. 3. Other anticoagulants, e.g., EDTA, can interfere with the creatinine enzymatic reaction and should not be used. Serum, rather than plasma, collection tubes should also be avoided as clotting activators can interfere with creatinine detection. 4. Solid wall black plates, e.g., Corning 3,915, should be used to avoid crosstalk between wells during measurements. 5. The fluorescent plate reader should have adjustable gain and be equipped with monochromators or filters to allow fluorescent measurements at Ex/Em of 495 nm/525 nm, 555 nm/ 585 nm, and 538 nm/587 nm for detection of fluorescein, rhodamine B, and the creatinine assay reaction product. 6. Calculate the amount of probe solution required for the mice (0.25 ml/mouse) and add 0.5 ml to determine the total probe stock volume that should be prepared. The extra 0.5 ml of probe stock solution will allow for a small aliquot to be retained for preparing standards as well as loss during filtration and gavage. 7. Cover the tube with foil to protect the stock solution from light. From this point on, tubes containing probe solution should always be covered with foil. 8. Fasting, gavage, retro-orbital blood collection, and probe use must be approved by the Institutional Animal Care and Use Committee (IACUC). Blood can also be collected from the tail vein. ä Fig. 6 (continued) (white), Na-K ATPase (green), and nuclei (blue) are shown for reference. Bar ¼ 20 μm. (c) Merged image showing early ulcer re-epithelialization (arrow) and reduced occludin (red) expression within damaged epithelium [54]. Phosphorylated MLC (white) is visible in endothelial cells within the ulcer bed (asterisk). Na-K ATPase (green) and nuclei (blue) are also shown. Bar ¼ 20 μm. (d) In the absence of damage, flux of all three probes is limited. (e) DSS-induced mucosal damage increases unrestricted pathway permeability, which can be measured as 70 kDa dextran flux. (f) Creatinine flux before and after DSS treatment. (g) 4 kDa dextran flux before and after DSS treatment. (h) 70 kDa dextran flux before and after DSS treatment. (i) The creatinine-to-70 kDa dextran ratio is increased, but, because 70 kDa dextran flux is increased, the fold-increase is proportionally smaller than that of creatinine. This indicates that creatinine is permeating via the unrestricted pathway; it crosses this pathway far more efficiently than the much larger 70 kDa dextran probe. (j) The 4 kDa dextran-to-70 kDa dextran ratio is increased nonsignificantly. This may have been statistically significant with a greater number of mice. The mean increase in the ratio is smaller than the mean increase in 4 kDa dextran flux. (k) The creatinine-to-4 kDa dextran ratio is unchanged, indicating that the increases in both creatinine and 4 kDa dextran are both due to unrestricted pathway upregulation. Mean  SD, **p < 0.01, *p < 0.05

264

Sandra D. Chanez-Paredes et al.

9. This method cannot be used to study mice with altered renal blood flow or clearance, as these will affect plasma probe concentrations. 10. If possible, a preliminary experiment should be performed at least 1 week prior to the actual experiment. Chow should be removed for 3 h, after which the mice are gavaged with water instead of probe. Chow can be returned to the cage 90 min later. Mice can be returned to normal housing conditions 3 h after gavage. This process acclimatizes the mice to handling, fasting, and gavage and will help minimize experimental noise due to stress [60]. 11. The proximal gut must be cleared of chow, bedding, and other materials to which probes can adsorb or be absorbed. Although mice are coprophagic, the amount of stool ingested over this short period is not problematic. A wire-bottom cage should not be used because it causes significant stress that can affect intestinal permeability [60]. 12. Staff must be trained in handling and gavage of mice. Improper techniques can induce stress that alters intestinal permeability and other physiological responses. 13. A 100 gavage needle will not reach the xyphoid process but is typically just long enough for use in adult C57BL/6 mice. Use of this needle prevents accidental over-insertion. Measuring the distance is critical if a longer needle or smaller mice are used. 14. For gavage, the head must be immobilized. To confirm proper restraint, hold a chow pellet next to the cheek. The mouse should not be able to turn its head to reach the pellet. Firm manual restraint reduces animal distress and also facilitates proper gavage technique. 15. The ball tip limits tissue damage and prevents accidental passage of the gavage needle into the trachea. Sedation or anesthesia should not be used, as these can increase the risk of aspiration pneumonia. 16. If the animal has difficulty breathing or there are any other signs of distress, the needle may be incorrectly placed. Withdraw and reposition the gavage needle before injecting the probe solution. 17. At least one healthy control mouse should be gavaged with water, without probes. If there is concern that the experimental model may affect basal plasma creatinine levels, separate wateronly control(s) should be generated for these mice. Similarly, when genetically modified mice are being studied, separate water-only controls should be created for each genotype until

Multiplex Intestinal Permeability Analysis

265

it is determined that basal plasma creatinine levels are unaffected by the genetic change(s). 18. Retro-orbital blood must be collected exactly 3 h after gavage. 19. Mice that fail to recover completely may have aspirated probes. This can happen due to gavage needle misplacement or regurgitation. These mice cannot be used for the assay, as they did not receive a full probe dose and may need to be sacrificed. 20. Because both gavage and phlebotomy require time, allow at least 3 min between mice and keep track of exactly when each mouse was gavaged. The minimal interval between mice should be determined by the time required for phlebotomy. 21. Because the tubes contain 50 μl of citrate buffer, which dilutes the plasma, it is important that the volume collected be consistent between mice. The fill line should be drawn between 175 μl and 250 μl total volume (but at exactly the same place on each tube), including the citrate buffer, depending on the size of the mice studied. 22. The anesthetic must be approved by the Institutional Animal Care and Use Committee and used under appropriate DEA permits. We typically induce anesthesia and analgesia using an i.p. injection containing ketamine (75 mg/kg) and xylazine (16 mg/kg). 23. The anesthetic dose should establish surgical plane anesthesia within ~8 min and maintain this until at least 15 min after administration. Surgical plane anesthesia can be assessed by the absence of reaction to toe or ear pinch. 24. If necessary, the small amount of blood that remains in the micropipette can be expelled into the collection tube using the bulb provided by the manufacturer. 25. A triple antibiotic ophthalmic ointment should be applied to the eye after phlebotomy. 26. Handle blood samples carefully to avoid hemolysis, which can release heme that fluoresces and interferes with measurements. 27. If the creatinine assay cannot be performed immediately, fluorescein and rhodamine fluorescences should be determined and plasma processed by dilution and spin column filtration before freezing. After filtration, samples can be stored at 80  C for up to 2 weeks and then thawed on ice before creatinine analysis. Because some creatinine degradation can occur, this approach should only be used if absolutely necessary. 28. The initial concentration of FITC-4 kDa dextran is 20 mM (80 mg/ml). Final concentrations of standards for fluorescein are 667 μM (1:30), 200 μM (1:100), 67 μM (1:300), 20 μM

266

Sandra D. Chanez-Paredes et al.

(1:1,000), 6.7 μM (1:3,000), 2 μM (1:10,000), 0.67 μM (1:30,000), 0.2 μM (1:100,000), 67 nM (1:300,000), 20 nM (1:1,000,000), and 7 nM (1:3,000,000). The initial concentration of rhodamine-70 kDa dextran is ~570 μM (40 mg/ml). Final concentrations of the standards are 19 μM (1:30), 5.7 μM (1:100), 1.9 μM (1:300), 570 nM (1:1,000), 190 nM (1:3,000), 57 nM (1:10,000), 19 nM (1:30,000), 5.7 nM (1:100,000), 1.9 nM (1:300,000), 0.57 nM (1:1,000,000), 0.19 nM (1:3,000,000). 29. Ideally, the standards should be diluted in plasma. However, the quantity of plasma needed would require exsanguination of more mice than is practical. Instead, a water blank is subtracted from the standards and a plasma blank from the mouse gavaged with water is subtracted from the plasma samples. 30. In our experience, the variance between technical replicates is minimal in these assays. In order to limit the amount of blood taken from each mouse, technical replicates are not included. In contrast, permeabilities of mice can vary significantly and biological replicates are essential. 31. The wavelengths and gains used for measurements may need to be optimized for the plate reader used. We use a Biotek Synergy HT with dual monochromators. Preliminary experiments should be performed to confirm the absence of bleed-through between channels. 32. Skip ahead to plasma dilution and filtration if the creatinine assay will not be performed immediately. The standards should be prepared fresh at time of assay. 33. The creatinine assay buffer is provided in the kit. It must be stored at 20  C but thawed and warmed to room temperature before use. To avoid multiple freeze-thaw cycles, store the assay buffer in aliquots equal to the volume needed for one experiment. 34. The creatinine assay buffer is very viscous. Take care to pipette from just below the meniscus to prevent buffer accumulation on the outside of the tip. Pipette slowly to avoid inaccurate measurements. 35. It is best to plan the layout of standards and samples in the plate before beginning. Because the standards can be very bright, it may be helpful to have an empty row or column between standards and samples. 36. The spin columns eliminate larger proteins, which can interfere with the creatinine assay. It is critical to process the samples only after fluorescent dextrans have been measured since 70 kDa dextran will not pass freely through the spin column filter.

Multiplex Intestinal Permeability Analysis

267

37. As noted above, technical replicates are not typically performed. New users may wish to verify consistency of technical replicates. 38. Prepare enough master mix for one extra reaction to allow for loss while pipetting the viscous solution. 39. New users may wish to prepare a small amount of master mix without creatininase (add 2 μl extra assay buffer instead) to assess background reactivity due to creatine or sarcosine within one or more samples. In our experience, this is not necessary routinely. 40. Cover the plate with foil to protect the fluorescent reaction product. 41. Although the creatinine assay reaction product fluorescence is qualitatively similar to rhodamine B, the creatinine assay reaction product intensity is far greater than that of any residual rhodamine B fluorescence that was not eliminated by the spin column. 42. RFU values for creatinine in probe-gavaged healthy mice are typically between 500 and 1500. RFU values for fluorescent dextran in probe-gavaged healthy mice typically exceed ~1000. The RFU of higher concentration standards may exceed the maximum measurable values for the gain settings selected as long as no experimental samples exceed maximum measurable values. 43. Because background has already been subtracted, the regression line should pass through the origin. The equation defining the relationship between fluorescent signal (x) and concentration ( y) is defined by y ¼ mx, where m represents the slope. If R2 is not 0.95 or greater, an error has been made, and the standard curve should be regenerated. Because creatinine standards are generated separately from the combined 4 kDa and 70 kDa dextran standards, errors should be present in the creatinine regression analysis only or both the fluorescein and rhodamine regression analyses. Problems with all three regression analyses indicate a global problem, such as plate reader malfunction or pipetting inaccuracy. 44. If more than one type of control, water-gavaged mouse is used, e.g., genetically-modified mice, the appropriate control should be used for each experimental sample, including healthy controls. 45. These values will be normalized in the next step. If raw values, i.e., actual probe concentrations, are needed, corrections for dilution with citrate buffer at time of blood collection and each subsequent dilution must be included. 46. Because normalized values are used, the healthy control group should have a mean of 1.0 for each ratio.

268

Sandra D. Chanez-Paredes et al.

Acknowledgments We thank Drs. Pei-Yun Tsai, Preeti Raju, Pawin Pongkorpsakol, Sam C. Nalle, Daniel R. Clayburgh, and other current and previous members of our laboratory for their contributions to development of these methods. We are indebted to Ms. Tiffany S. Davanzo (Slaybaugh Studios) for her beautiful illustrations and Ms. Heather Marlett (Nationwide Histology) for her outstanding tissue preparation. This work was supported by NIH grants to JRT (R01DK61931, R01DK68271, R24DK099803) and the Harvard Digestive Disease Center (P30DK034854). Conflicts of Interest: JRT is a cofounder of and hold shares in Thelium Therapeutics. References 1. Turner JR (2009) Intestinal mucosal barrier function in health and disease. Nat Rev Immunol 9:799–809. https://doi.org/10.1038/ nri2653 2. Peterson LW, Artis D (2014) Intestinal epithelial cells: regulators of barrier function and immune homeostasis. Nat Rev Immunol 14:141–153. https://doi.org/10.1038/ nri3608 3. McDole JR, Wheeler LW, McDonald KG et al (2012) Goblet cells deliver luminal antigen to CD103+ dendritic cells in the small intestine. Nature 483:345–349. https://doi.org/10. 1038/nature10863 4. Niess JH, Brand S, Gu X et al (2005) CX3CR1-mediated dendritic cell access to the intestinal lumen and bacterial clearance. Science 307:254–258. https://doi.org/10. 1126/science.1102901 5. Lai NY, Musser MA, Pinho-Ribeiro FA et al (2020) Gut-innervating nociceptor neurons regulate Peyer’s patch microfold cells and SFB levels to mediate salmonella host defense. Cell 180(33-49):e22. https://doi.org/10.1016/j. cell.2019.11.014 6. Farquhar M, Palade G (1963) Junctional complexes in various epithelia. J Cell Biol 17:375–412 7. Kottra G, Fromter E (1983) Functional properties of the paracellular pathway in some leaky epithelia. J Exp Biol 106:217–229 8. Furuse M, Hata M, Furuse K et al (2002) Claudin-based tight junctions are crucial for the mammalian epidermal barrier: a lesson from claudin-1-deficient mice. J Cell Biol 156:1099–1111. https://doi.org/10.1083/ jcb.200110122

9. Umeda K, Ikenouchi J, Katahira-Tayama S et al (2006) ZO-1 and ZO-2 independently determine where claudins are polymerized in tightjunction strand formation. Cell 126:741–754. https://doi.org/10.1016/j.cell.2006.06.043 10. Otani T, Nguyen TP, Tokuda S et al (2019) Claudins and JAM-A coordinately regulate tight junction formation and epithelial polarity. J Cell Biol 218:3372–3396. https://doi.org/ 10.1083/jcb.201812157 11. Nelms B, Dalomba NF, Lencer W (2017) A targeted RNAi screen identifies factors affecting diverse stages of receptor-mediated transcytosis. J Cell Biol 216:511–525. https://doi. org/10.1083/jcb.201609035 12. Ko SY, Pegu A, Rudicell RS et al (2014) Enhanced neonatal Fc receptor function improves protection against primate SHIV infection. Nature 514:642–645. https://doi. org/10.1038/nature13612 13. Amasheh S, Meiri N, Gitter AH et al (2002) Claudin-2 expression induces cation-selective channels in tight junctions of epithelial cells. J Cell Sci 115:4969–4976. https://doi.org/10. 1242/jcs.00165 14. Van Itallie CM, Fanning AS, Anderson JM (2003) Reversal of charge selectivity in cation or anion-selective epithelial lines by expression of different claudins. Am J Physiol Renal Physiol 285:F1078–F1084. https://doi.org/ 10.1152/ajprenal.00116.2003 15. Weber CR, Liang GH, Wang Y et al (2015) Claudin-2-dependent paracellular channels are dynamically gated. elife 4:e09906. https://doi. org/10.7554/eLife.09906 16. Li J, Zhuo M, Pei L et al (2014) Comprehensive cysteine-scanning mutagenesis reveals

Multiplex Intestinal Permeability Analysis claudin-2 pore-lining residues with different intrapore locations. J Biol Chem 289:6475–6484. https://doi.org/10.1074/ jbc.M113.536888 17. Wada M, Tamura A, Takahashi N et al (2013) Loss of claudins 2 and 15 from mice causes defects in paracellular Na+ flow and nutrient transport in gut and leads to death from malnutrition. Gastroenterology 144:369–380. https://doi.org/10.1053/j.gastro.2012.10. 035 18. Anderson JM, Van Itallie CM (2009) Physiology and function of the tight junction. Cold Spring Harb Perspect Biol 1:a002584. https:// doi.org/10.1101/cshperspect.a002584 19. Colegio OR, Van Itallie C, Rahner C et al (2003) Claudin extracellular domains determine paracellular charge selectivity and resistance but not tight junction fibril architecture. Am J Physiol Cell Physiol 284:C1346–C1354. https://doi.org/10.1152/ajpcell.00547.2002 20. Colegio OR, Van Itallie CM, McCrea HJ et al (2002) Claudins create charge-selective channels in the paracellular pathway between epithelial cells. Am J Physiol Cell Physiol 283: C142–C147. https://doi.org/10.1152/ ajpcell.00038.2002 21. Van Itallie CM, Holmes J, Bridges A et al (2008) The density of small tight junction pores varies among cell types and is increased by expression of claudin-2. J Cell Sci 121:298–305. https://doi.org/10.1242/jcs. 021485 22. Angelow S, Yu AS (2009) Structure-function studies of claudin extracellular domains by cysteine-scanning mutagenesis. J Biol Chem 284:29205–29217. https://doi.org/10. 1074/jbc.M109.043752 23. Tanaka H, Yamamoto Y, Kashihara H et al (2016) Claudin-21 has a paracellular channel role at tight junctions. Mol Cell Biol 36:954–964. https://doi.org/10.1128/ MCB.00758-15 24. Rosenthal R, Gunzel D, Piontek J et al (2020) Claudin-15 forms a water channel through the tight junction with distinct function compared to claudin-2. Acta Physiol (Oxf) 228:e13334. https://doi.org/10.1111/apha.13334 25. Holmes JL, Van Itallie CM, Rasmussen JE et al (2006) Claudin profiling in the mouse during postnatal intestinal development and along the gastrointestinal tract reveals complex expression patterns. Gene Expr Patterns 6:581–588. https://doi.org/10.1016/j.modgep.2005.12. 001 26. Ong M, Yeruva S, Sailer A et al (2020) Differential regulation of claudin-2 and claudin-15

269

expression in children and adults with malabsorptive disease. Lab Investig 100:483–490. https://doi.org/10.1038/s41374-019-03248 27. Heller F, Florian P, Bojarski C et al (2005) Interleukin-13 is the key effector Th2 cytokine in ulcerative colitis that affects epithelial tight junctions, apoptosis, and cell restitution. Gastroenterology 129:550–564. https://doi.org/ 10.1016/j.gastro.2005.05.002 28. Tsai PY, Zhang B, He WQ et al (2017) IL-22 upregulates epithelial claudin-2 to drive diarrhea and enteric pathogen clearance. Cell Host Microbe 21:671–681.e4. https://doi.org/10. 1016/j.chom.2017.05.009 29. Luettig J, Rosenthal R, Barmeyer C et al (2015) Claudin-2 as a mediator of leaky gut barrier during intestinal inflammation. Tissue Barriers 3:e977176. https://doi.org/10. 4161/21688370.2014.977176 30. Wu RL, Vazquez-Roque MI, Carlson P et al (2017) Gluten-induced symptoms in diarrheapredominant irritable bowel syndrome are associated with increased myosin light chain kinase activity and claudin-15 expression. Lab Investig 97:14–23. https://doi.org/10.1038/ labinvest.2016.118 31. Weber CR, Raleigh DR, Su L et al (2010) Epithelial myosin light chain kinase activation induces mucosal interleukin-13 expression to alter tight junction ion selectivity. J Biol Chem 285:12037–12046. https://doi.org/ 10.1074/jbc.M109.064808 32. Raju P, Shashikanth N, Tsai PY et al (2020) Inactivation of paracellular cation-selective claudin-2 channels attenuates immunemediated experimental colitis in mice. J Clin Invest 130:5197–5208. https://doi.org/10. 1172/JCI138697 33. Marchiando AM, Shen L, Graham WV et al (2010) Caveolin-1-dependent occludin endocytosis is required for TNF-induced tight junction regulation in vivo. J Cell Biol 189:111–126. https://doi.org/10.1083/jcb. 200902153 34. Clayburgh DR, Barrett TA, Tang Y et al (2005) Epithelial myosin light chain kinase-dependent barrier dysfunction mediates T cell activationinduced diarrhea in vivo. J Clin Invest 115:2702–2715. https://doi.org/10.1172/ JCI24970 35. Zolotarevsky Y, Hecht G, Koutsouris A et al (2002) A membrane-permeant peptide that inhibits MLC kinase restores barrier function in in vitro models of intestinal disease. Gastroenterology 123:163–172. https://doi.org/10. 1053/gast.2002.34235

270

Sandra D. Chanez-Paredes et al.

36. Wang F, Graham WV, Wang Y et al (2005) Interferon-gamma and tumor necrosis factoralpha synergize to induce intestinal epithelial barrier dysfunction by up-regulating myosin light chain kinase expression. Am J Pathol 166:409–419. https://doi.org/10.1016/ s0002-9440(10)62264-x 37. Su L, Nalle SC, Shen L et al (2013) TNFR2 activates MLCK-dependent tight junction dysregulation to cause apoptosis-mediated barrier loss and experimental colitis. Gastroenterology 145:407–415. https://doi.org/10.1053/j. gastro.2013.04.011 38. Purandare S, Offenbartl K, Westrom B et al (1989) Increased gut permeability to fluorescein isothiocyanate-dextran after total parenteral nutrition in the rat. Scand J Gastroenterol 24:678–682. https://doi.org/ 10.3109/00365528909093108 39. Dawson PA, Huxley S, Gardiner B et al (2009) Impaired intestinal function in the hyposulphataemic NaS1 null mouse. Gut 58:910–919. https://doi.org/10.1136/gut.2007.147595 40. Novosad VL, Richards JL, Phillips NA et al (2013) Regional susceptibility to stressinduced intestinal injury in the mouse. Am J Physiol Gastrointest Liver Physiol 305: G418–G426. https://doi.org/10.1152/ ajpgi.00166.2013 41. Roxas JL, Koutsouris A, Bellmeyer A et al (2010) Enterohemorrhagic E. coli alters murine intestinal epithelial tight junction protein expression and barrier function in a Shiga toxin independent manner. Lab Invest 90 (8):1152–1168. https://doi.org/10.1038/ labinvest.2010.91 42. Gerlach K, Hwang Y, Nikolaev A et al (2014) TH9 cells that express the transcription factor PU.1 drive T cell-mediated colitis via IL-9 receptor signaling in intestinal epithelial cells. Nat Immunol 15:676–686. https://doi.org/ 10.1038/ni.2920 43. Rawat M, Nighot M, Al-Sadi R et al (2020) IL1B increases intestinal tight junction permeability by up-regulation of MIR200C-3p, which degrades occludin mRNA. Gastroenterology 159:1375–1389. https://doi.org/10. 1053/j.gastro.2020.06.038 44. Meisel M, Hinterleitner R, Pacis A et al (2018) Microbial signals drive pre-leukaemic myeloproliferation in a Tet2-deficient host. Nature 557:580–584. https://doi.org/10.1038/ s41586-018-0125-z 45. Nalle SC, Zuo L, Ong M et al (2019) Graftversus-host disease propagation depends on increased intestinal epithelial tight junction permeability. J Clin Invest 129:902–914. https://doi.org/10.1172/JCI98554

46. Bird L (2014) T cells: IL-9 breaks down barriers. Nat Rev Immunol 14:432. https://doi. org/10.1038/nri3709 47. Kissoon-Singh V, Moreau F, Trusevych E et al (2013) Entamoeba histolytica exacerbates epithelial tight junction permeability and proinflammatory responses in Muc2(/) mice. Am J Pathol 182:852–865. https://doi.org/ 10.1016/j.ajpath.2012.11.035 48. Zhang YG, Lu R, Xia Y et al (2018) Lack of vitamin D receptor leads to hyperfunction of claudin-2 in intestinal inflammatory responses. Inflamm Bowel Dis 25(1):97. https://doi. org/10.1093/ibd/izy292 49. Ahmad R, Chaturvedi R, Olivares-Villagomez D et al (2014) Targeted colonic claudin-2expression renders resistance to epithelial injury, induces immune suppression, and protects from colitis. Mucosal Immunol 7:1340–1353. https://doi.org/10.1038/mi. 2014.21 50. Pongkorpsakol P, Turner JR, Zuo L (2020) Culture of intestinal epithelial cell monolayers and their use in multiplex macromolecular permeability assays for in vitro analysis of tight junction size selectivity. Curr Protoc Immunol 131:e112. https://doi.org/10.1002/cpim. 112 51. Fihn BM, Sjoqvist A, Jodal M (2000) Permeability of the rat small intestinal epithelium along the villus-crypt axis: effects of glucose transport. Gastroenterology 119:1029–1036. https://doi.org/10.1053/gast.2000.18148 52. Pappenheimer JR, Renkin EM, Borrero LM (1951) Filtration, diffusion and molecular sieving through peripheral capillary membranes; a contribution to the pore theory of capillary permeability. Am J Phys 167:13–46. https:// doi.org/10.1152/ajplegacy.1951.167.1.13 53. Bassingthwaighte JB (2006) A practical extension of hydrodynamic theory of porous transport for hydrophilic solutes. Microcirculation 13:111–118. https://doi.org/10.1080/ 10739680500466384 54. Kuo WT, Shen L, Zuo L et al (2019) Inflammation-induced occludin downregulation limits epithelial apoptosis by suppressing caspase-3 expression. Gastroenterology 157:1323–1337. https://doi.org/10.1053/j. gastro.2019.07.058 55. Menzies IS, Laker MF, Pounder R et al (1979) Abnormal intestinal permeability to sugars in villous atrophy. Lancet 2:1107–1109. https:// doi.org/10.1016/s0140-6736(79)92507-8 56. Arrieta MC, Bistritz L, Meddings JB (2006) Alterations in intestinal permeability. Gut

Multiplex Intestinal Permeability Analysis 55:1512–1520. https://doi.org/10.1136/ gut.2005.085373 57. Hollander D (1992) The intestinal permeability barrier. A hypothesis as to its regulation and involvement in Crohn’s disease. Scand J Gastroenterol 27:721–726. https://doi.org/10. 3109/00365529209011172 58. Folin O, Morris JL (1914) On the determination of creatinine and creatine in urine. J Biol Chem 17:469–473. https://doi.org/10. 1016/s0021-9258(18)88386-7

271

59. den Elzen WPJ, Cobbaert CM, Klein Gunnewiek JMT et al (2018) Glucose and total protein: unacceptable interference on Jaffe creatinine assays in patients. Clin Chem Lab Med 56:e185–e187. https://doi.org/10. 1515/cclm-2017-1170 60. Meddings JB, Swain MG (2000) Environmental stress-induced gastrointestinal permeability is mediated by endogenous glucocorticoids in the rat. Gastroenterology 119:1019–1028. https://doi.org/10.1053/gast.2000.18152

Methods in Molecular Biology (2021) 2367: 273–290 DOI 10.1007/7651_2021_347 © Springer Science+Business Media, LLC 2021 Published online: 17 April 2021

Establishment of Intestinal Epithelial Cell Monolayers and Their Use in Calcium Switch Assay for Assessment of Intestinal Tight Junction Assembly Pawin Pongkorpsakol, Wilasinee Satianrapapong, Preedajit Wongkrasant, Peter R. Steinhagen, Nuttha Tuangkijkul, Nutthapoom Pathomthongtaweechai, and Chatchai Muanprasat Abstract Intestinal barrier function relies primarily on the assembly and integrity of tight junctions, which forms a size-selective barrier. This barrier restricts paracellular movement of solutes in various types of epithelia. Of note, extracellular Ca2+ concentration affects tight junction assembly. Therefore, the removal and re-addition of Ca2+ into cell culture medium of cultured intestinal epithelial cells causes destabilization and reassembly of tight junction to membrane periphery near apical surface, respectively. Based on this principle, the Ca2+-switch assay was established to investigate tight junction assembly in fully differentiated intestinal epithelial cells. This chapter provides a stepwise protocol for culture of intestinal epithelial cell monolayers using T84 cell line as an in vitro model and the Ca2+-switch assay for evaluating tight junction assembly. Key words Ca2+-switch assay, Paracellular permeability, Barrier function, Tight junction, Transepithelial electrical resistance (TER), Epithelial cells

1

Introduction One of the critical roles of the intestinal epithelium is to establish a tissue barrier, which tightly controls the interactions between the gut microbiota and host internal milieu. The intestinal epithelial barrier has been considered as a part of the innate immune system. For example, this barrier protects against invasion of pathogens residing the gut lumen. Moreover, intestinal epithelial barrier function establishes polarity of intestinal epithelial cells, which also supports vectorial transport of nutrient and fluid absorption, as well as eliminates waste products [1–4]. Intestinal epithelial barrier function depends mainly on assembly of tight junction proteins at the peri-junctional actomyosin ring (PAMR) near the apical membrane. Importantly, compromised intestinal barrier function caused by tight junction disassembly is associated with pathogenesis and progression of many diseases, not only intestinal, but also systemic

273

274

Pawin Pongkorpsakol et al.

diseases, i.e., inflammatory bowel diseases (IBD) [5, 6] and graftversus-host disease (GVHD) [7, 8]. Currently, there are no FDA-approved drugs for restoring the epithelial barrier [3]. To achieve this goal, a comprehensive understanding of both disease pathogenesis and mechanistic insights into tight junction barrier regulation need to be reached. Based on its ultrastructural morphology, the epithelial tight junction was thought to be static. However, fluorescence recovery after photobleaching (FRAP) analysis of EGFP-tagged tight junction proteins expressed in Madin-Darby Canine Kidney (MDCK) monolayers revealed that, in addition to static claudin-1, ZO-1, and occludin are highly dynamic at steady state [9]. Indeed, occludin passively diffuses within the tight junction and along the lateral membrane. On the other hand, ZO-1 can diffuse to the cytoplasmic site in an energy-dependent manner [9]. Apart from steady state, tight junction proteins can be endocytosed in response to some inflammatory stimuli. In physiological condition, tight junction permeability can be increased in parallel with activation of Na+glucose cotransport [10, 11], this is to supplement the saturated active transcellular nutrient absorption [12–15]. In pathogenic condition, TNF-α and LIGHT can increase tight junction permeability via myosin light-chain kinase (MLCK)-dependent mechanism [16–20], and this event occurs in IBD. Moreover, lipopolysaccharides (LPS) suppressed the trafficking process of tight junction proteins and delayed tight junction assembly [21]. Therefore, trafficking and assembly of tight junction proteins are key factors that determine the integrity of tight junction at the intercellular space. Tight junction protein expression, formation, and assembly are essential for survival of multicellular organisms [22–25]. Evaluation of tight junction integrity can be simply performed by measuring the transepithelial electrical resistance (TER) of epithelial cell monolayers, which mainly reflects paracellular resistance, approximately 75–94% of the total passive ion flux across intestinal epithelia [26–32], together with immunofluorescence staining of tight junction protein and FITC-dextran flux assay [31, 33]. High-resolution analyses of tight junction permeability can be done by multiplex macromolecular permeability [34] and bi-ionic potential assays [35, 36] for in vitro assessment of tight junction size and charge selectivity, respectively. Several lines of evidence indicated that tight junction assembly in various cell types requires optimal extracellular Ca2+ concentration [37–43]. In fact, extracellular Ca2+ directly interacts and stabilizes tight junction integrity [38, 40]. However, the process of tight junction barrier re-assembly also depends on intracellular Ca2+ signaling [37]. Therefore, depletion and supplementation of Ca2+ into cell culture medium of differentiated epithelial cells can lead to destabilization and re-assembly of tight junction proteins,

Establishment of Intestinal Epithelial Cell Monolayers and Their Use in. . .

275

Fig. 1 Principle of the Ca2+-switch assay. Cell culture medium containing Ca2+ allows assembly of tight junctional proteins (left). Removal of Ca2+ results in tight junction disassembly and increased flux of molecules across the paracellular route (center). Re-addition of Ca2+ to cell culture medium induces reassembly of tight junctions and restores barrier function (right)

respectively. This method is termed Ca2+-switch assay. This method is used to investigate tight junction assembly in differentiated epithelial cell monolayers [21, 31, 44]. This simple and useful in vitro assay mimics the event of tight junction disruption, and to discover new drug candidates for the restoration of tight junction integrity. For example, this assay has been used in recent studies to identify AMP-activated protein kinase (AMPK) and zinc-sensing G-protein coupled receptor 39 (GPR39) as drug targets to modify paracellular barrier function of tight junction [31, 44]. Of particular interest, using the Ca2+-switch assay, butyrate, and fructo-oligosaccharides have recently been demonstrated as the activators of tight junction assembly [21, 45]. This chapter illustrates how to culture intestinal epithelial cell monolayers using T84 cells as an in vitro model and how to perform Ca2+-switch assay in order to investigate tight junction assembly (Fig. 1).

2

Materials The Ca2+-switch assay can be applied to various types of epithelial cells to determine tight junction assembly in fully differentiated cell monolayers including T84, Caco-2BBe, or MDCK cell lines. Here, we provide example experiments using T84 cells grown as monolayers on Transwell semipermeable supports. Assembly and disassembly of tight junctions are easily determined by TER measurements (Fig. 2), together with immunofluorescence staining of tight junction proteins, and/or FITC-dextran (4 kDa) permeability assay. Cell culture medium and buffers used for this method should be prepared in ultrapure (deionized) water (see Note 1).

276

Pawin Pongkorpsakol et al.

Fig. 2 Setting for TER measurement. A chopstick electrode is attached to the Epithelial Volt/ohm meter (EVOM) and measures TER of an epithelial cell monolayer grown on Transwell inserts. Circuit of several factors that influence resistance of a cell monolayer 2.1

Reagents Needed

1. Gibco® S-MEM (Spinner Modification, MEM for suspension cultures), no calcium, no glutamine (Thermo Fisher Scientific, Catalog number: 11380-037). This type of culture medium is used for induction of tight junction disassembly in fully differentiated intestinal epithelial cell monolayers. 2. Fetal bovine serum (Thermo Fisher Scientific, Catalog number: 10270-106). 3. 0.25% Trypsin-EDTA (Thermo Fisher Scientific, Catalog number: 25200-056). 4. Collagen type I, rat tail (Corning, Catalog number: 354236, in 0.02N acetic acid to prevent polymerization) (see Notes 2 and 3). 5. HEPES (1 M) solution (Sigma-Aldrich, catalog number: H0887). 6. Hanks’ Balanced Salt solution (HBSS), powders, without phenol red (Sigma-Aldrich, catalog number: H1387). This solution can be manually prepared, based on supplier recipes: NaCl (8 g/L), Na2HPO4 (0.04788 g/L), MgSO4 (0.09767 g/L), KCl (0.4 g/L), KH2PO4 (0.06 g/L), CaCl2 (0.1396 g/L), D-glucose (1 g/L). Dissolve HBSS compound in ~750–800 mL ultrapure water. Buffers HBSS with 15 mM HEPES (Add 15 mL 1 M HEPES into 1 L HBSS). Stir the

Establishment of Intestinal Epithelial Cell Monolayers and Their Use in. . .

277

solution until all compounds are disappeared. Measure and adjust the solution pH to 7.4 (see Note 4). Adjust the volume to 1 L. Sterilize HBSS solution by filtering through 0.2-μm bottle-top filter into pre-autoclaved bottle. 7. 70% (v/v) ethanol, nondenatured. 8. Culture medium appropriate for T84 cell line: DMEM/F-12 (Dulbecco’s Modified Eagle Medium/Nutrient Mixture F-12; powder), with HEPES, L-glutamine, and Phenol Red, without Na2HPO4 (Thermo Fisher Scientific, Catalog number: 12400024). Supplement with 10% fetal bovine serum and add Na2HPO4 (1.2 g/L). Adjust the medium pH to 7.4 (see Note 4). Sterilize DMEM/F-12 with supplement using 0.2-μm bottle-top filter. 9. 4 kDa Fluorescein isothiocyanate-dextran (FITC-dextran; Sigma-Aldrich, catalog number: 46944). 10. ZO-1 Monoclonal Antibody (ZO1-1A12) (Thermo Fisher Scientific, Catalog number: 33-9100). 2.2

Materials Needed

1. T84 cell line (ATCC® CCL-248™), human colorectal carcinoma cell line, derived from a lung metastatic site in a 72-yearold male. 2. T75 flasks, culture area: 75 cm2 (Corning, catalog number: 07-202-000). 3. Polyester Membrane Insert, Sterile, 12-well format, 12-mm Transwell® with Pore size 0.4 μm, membrane thickness 10 μm, cell growth area 1.12 cm2 (Corning, catalog number: 3460). 4. 12-well Clear TC-treated Multiple Well Plates (Corning, catalog number: 3512). 5. 96-well Black Flat Bottom Polystyrene Not Treated Microplate (Corning, catalog number: 3915). 6. Pre-autoclaved bottles (500 mL). 7. Pre-autoclaved microcentrifuge tubes (0.5 mL, 1.5 mL).

2.3

Equipment

1. Fluorescence microplate reader. 2. Epithelial Volt/Ohm Meter (EVOM). 3. CO2 Incubator. 4. Water bath. 5. Warming plate. 6. Electronic multi-dispensing pipette. 7. Pre-autoclaved beakers.

278

2.4

3

Pawin Pongkorpsakol et al.

Cell Culture

For maintenance, T84 cells should be cultured in a T75 flask. Cell culture medium should be renewed every two days. Confluent T84 cells should be split when cells show 80% confluence. The cells in a T75 flask should be trypsinized once a week. Dilute T84 cells 1:10 for passage. For experimental use, T84 cells are cultured as monolayers on collagen-coated Polyester Membrane Transwell Inserts (Corning). Following 8–12 days after seeding, T84 cells are ready for experiments. Overgrown T84 cell monolayers should not be used. In addition, monolayers cultured for less than 7 days and for more than 12 days are not recommended for experimental use. Based on our experience, optimal experimental conditions for performing a Ca2+-switch assay can be achieved with monolayers following 10 days after seeding.

Methods This protocol is optimized for T84 cells grown as monolayers in Transwell inserts (1.12 cm2) only. In case of larger or smaller sizes of Transwell inserts, or other cell types, this protocol needs to be re-optimized for the amount of cells per Transwell insert, time period of cell culture before experiment and in S-MEM medium. In this part, we provide a detailed protocol for the maintenance of T84 cell cultures, growth of T84 cell monolayers and Ca2+-switch assay. We also show how to perform TER measurements, FITC dextran flux assay, immunofluorescence staining and provide analysis and interpretation of our experimental data.

3.1 Establishment of T84 Cell Monolayers

1. Coat Transwell inserts with rat tail Collagen I solution and let it dry overnight in a laminar flow hood (see Note 5). 2. Wash the apical chamber of each Transwell insert with 2 mL sterile HBSS to eliminate residual acetic acid. 3. Gently remove most of the HBSS from both the apical and basal chambers, take care and do not touch the insert membrane in order to prevent membrane damage and its collagen coating. 4. When the T84 cell culture in the T75 flask reaches ~80% confluency, remove the culture medium completely. We highly recommend the use of 80% confluent T84 cells, as this is optimal for cell plating. Do not use T84 cells with very low or very high confluencies (Fig. 3). 5. Warm culture medium and pre-warm 1 PBS in a water bath, set to 37  C. 6. Rinse T84 cells in T75 flask once with 10 mL sterile/filtered pre-warm 1 PBS and incubate T84 cell in 10 mL sterile/ filtered 1 pre-warmed PBS for 20 min at 37  C, 5% CO2.

Establishment of Intestinal Epithelial Cell Monolayers and Their Use in. . .

279

Fig. 3 T84 cell morphology and density. Following trypsinization and splitting of T84 cells in a T75 flask, T84 cells are mainly organized and reaches ~30% confluence within few days (A). During growth phase (Days 7–9), T84 cells grown in a T75 flask become confluent and reach a confluency of ~80% (B), and 90% (C) in T75 flasks

7. Use a light microscope to check the morphology of the T84 cells after 20 min of PBS incubation, this should result in a starting/slight dissociation of neighboring cells. 8. Entirely remove PBS from the T75 flask. 9. To trypsinize T84 cells, add 2 mL of 0.25% Trypsin/EDTA into the T75 flask containing T84 cells and incubate for 10 min at 37  C, 5% CO2 (see Note 6). 10. Add 8 mL pre-warmed culture medium (DMEM/F-12) into the T75 flask to inactivate trypsin/EDTA. 11. Resuspend T84 cells by pipetting up and down for at least 10 times to dissociate cells completely. 12. Use a light microscope to check the morphology of T84 cells. T84 cells should either be a clump of three to five cells or single cells. If this is not the case, pipet five more times up and down and check the morphology of the T84 cell suspension again (see Note 7). 13. Determine the total amount of T84 cell and dilute the suspended T84 cells with culture medium to obtain the working density of 106 cells/mL (see Note 8). 14. Seed 0.5 mL of the diluted T84 cell suspension into each Transwell insert (12-well format) and add 1.5 mL culture medium into each basal chamber of the plate. 15. Change culture media every two days, for example, on Monday, Wednesday, and Friday.

280

Pawin Pongkorpsakol et al.

Fig. 4 Time-dependent TER development of T84 cell monolayers. Following cell culture, T84 cells were seeded onto Transwell insert membranes and grown as monolayers. While TER values are low during the first days, T84 cell monolayers develop higher TER values after day 6

16. For media renewal, remove the old culture medium only from the basal chamber, do not aspirate medium from the apical chamber to avoid monolayer damage. 17. Carefully add 2 mL fresh DMEM/F-12 medium into the apical chamber of the Transwell inserts. Let the fresh cell culture medium overflow the apical chamber as this will add fresh cell culture medium to the basal chamber as well (see Notes 9–11). 18. Culture T84 cell monolayers in Transwell membrane inserts for 8–12 days, T84 cell monolayers will be ready to use and TER should be ~3000–4000 Ω cm2 (see Notes 11 and 12). We determined TER by using the EVOM and chopstick electrodes. Example of data: Based on our observation, TER of the T84 cell monolayers will usually develop at day 6 after plating (TER ~500–900 Ω cm2), and progressively increases and reaches TER values of ~2000–4000 Ω cm2 around days 8–10 following plating (Fig. 4). Based on our experience, 10-dayold T84 cell monolayers are suitable for this experiment. 3.2 Ca2+Switch Assay

1. At day 10 post-plating, measure TER to make sure that T84 cell monolayers are ready for the experiment (see Notes 11 and 12). If TER values of T84 cell monolayers are still below the recommended TER of ~2000–4000 Ω cm2, do not use monolayers and continue T84 cell monolayer culture for a few more days with regular renewal of cell culture medium (see Notes 13 and 14).

Establishment of Intestinal Epithelial Cell Monolayers and Their Use in. . .

281

Fig. 5 Effect of S-MEM medium on %TER of T84 monolayers and 4 kDa FITC-dextran permeability assay. Ca2+-free S-MEM medium results in a time-dependent TER drop (A). 4 kDa FITC-Dextran permeability increases in T84 monolayers incubated in Ca2+-free S-MEM for 16 h (B)

2. If the T84 cell monolayers reach the recommended TER values, gently remove cell culture medium. Make sure to keep the monolayer intact and start with removal of basal cell culture medium followed by apical cell culture medium of Transwell inserts (see Note 15). 3. Wash the T84 monolayers once with S-MEM medium, pre-warmed in a water bath set to 37  C, in order to remove regular culture medium (DMEM/F-12) containing Ca2+ as much as possible (see Note 15). 4. Start with adding 0.5 mL of S-MEM medium to the apical chamber followed by adding 1.5 mL of S-MEM medium to the basal chamber. Incubate T84 cell monolayers with S-MEM medium for 16 h at 37  C in a CO2 incubator (see Notes 16 and 17). Example of data: Time-dependent effect of S-MEM medium on TER in T84 cell monolayers (Fig. 5). (a) After incubation with S-MEM medium for 2 h, TER of T84 drops ~25% and slightly decrease. It reaches a 50% drop at 16 h after culturing the cell monolayers with S-MEM medium. At 24 h, TER decreases ~>90% from basal level, this is very extreme to be rescued (Fig. 5A). (b) At 16 h, permeability of 4 kDa FITC-dextran is significantly increased in S-MEM-cultured T84 cell monolayers compared to normal medium (DMEM/F-12) (see Note 18), indicating that S-MEM medium increases, sizeselective, tight junction permeability (Fig. 5B).

282

Pawin Pongkorpsakol et al.

Fig. 6 TER recovery after Ca2+ re-addition. Ca2+ removal results in a significant TER drop of T84 cell monolayers. Following Ca2+ re-addition, TER increases within a few hours and nearly reaches baseline TER

5. At 16 h after incubating the T84 cell monolayers with S-MEM medium, these monolayers that will drop ~50% can be used for the tight junction assembly experiment. Gently aspirate S-MEM medium from both the apical and basal chamber of the Transwells. 6. For control group, T84 monolayers are cultured with regular medium (DMEM/F-12) containing Ca2+ with vehicle (e.g., DMSO). This leads to recovery of tight junction barrier function again with slow kinetic rate. At 6 h, TER of T84 cell monolayers will be recovered ~90–100% (Fig. 6). 7. For treatment group, T84 monolayers are treated with test compounds to investigate its effect on tight junction assembly. Here, we show an example data using a GPR39 agonist (TC-G 1008; 10 μM), to enhance tight junction assembly (Fig. 7). TER measurements are performed at various time points. Immunofluorescence staining of ZO-1 can be performed to confirm the localization of tight junction marker for each condition (see Note 19).

4

Notes 1. Ultrapure or deionized water is processed through purification cartridges to reach a sensitivity of 18 MΩ cm (at 25  C). This type of water is highly recommended to use for the experiment to avoid possible sources of contamination. 2. Collagen coating is one of the most important steps. Then, cells may not be grown properly if coating process is not

Establishment of Intestinal Epithelial Cell Monolayers and Their Use in. . .

283

Fig. 7 Effects of GPR39 agonist on tight junction assembly. TER drops following Ca2+ depletion (A). Treatment with G protein-coupled receptor 39 (GPR39) agonist results in restoration of TER with higher values compared to baseline after 2 h. In contrast, vehicle treatment results in low TER values (A). Confluent T84 cell monolayer (B) and T84 monolayer disruption following Ca2+ depletion (C). Re-addition of Ca2+ and treatment with vehicle in the control group (D) and an agonist of G-protein coupled receptor 39 (GPR39) (E)

optimal. We recommend using 3 μg/cm2 Collagen I Rat Tail for T84 cells grown in 12-well format Transwell insert (Corning, catalog number: 3460). Dilution of Collagen I Rat Tail can be done in 70% EtOH (Molecular grade, 200 proof). 3. Collagen I Rat Tail at concentration of 3 μg/cm2 is recommended for coating Transwell insert for T84 cells. Since we use 1.12 cm2 Transwell insert, we therefore add 1 μg Collagen I Rat Tail into each Transwell insert. Optimal volume of 1 μg Collagen I Rat Tail is 150 μL/insert. The electronic multidispensing pipette is recommended for this coating step, which would accurately provide equal amount of Collagen coating solution into each Transwell insert and eliminate some possible human errors. Of note, different cell types and Transwell inserts may require different coating conditions. Therefore, concentration and volume of Collagen I Rat Tail need to be optimized before starting the experiment. 4. The pH of culture medium (DMEM/F-12) and HBSS buffer should be carefully checked and adjusted to 7.4 because there is evidence reporting that tight junction integrity would be interfered by changing extracellular pH [46]. 5. Dilute Collagen I Rat Tail in 70% ethanol (nondenatured) to obtain a stock concentration of 20 μg/mL. Add 150 μL of Collagen I Rat Tail (20 μg/mL) onto each Transwell insert to

284

Pawin Pongkorpsakol et al.

achieve a concentration of 3 μg/cm2. Replace the lid of the Transwell plates in which the plate is partially open to the air but still covered to protect against contamination. Leave collagen-coated Transwell plates in a laminar flow hood for 6 h as minimum, or overnight to allow the ethanol to evaporate. In our experience, overnight coating results in optimal conditions. 6. Confluent and post-confluent conditions of T84 cells are not recommended for all culture purposes because T84 cells will undergo differentiation causing inconsistent results. 7. As mentioned above, after trypsinizing, most of the T84 cells in T75 flask should be dispersed individually or forming a clump of 3–5 cells. If larger clusters of the cells still remain, the T84 cell suspension should be mixed by pipetting up and down again for five more times. In some circumstances, larger clumps may persist. This may be due to over-confluence. In this case, we do not recommend to continue mixing the cells by pipetting because it can lead to cell damage, however, seeding large cell clusters may not result in a uniform morphology of T84 monolayers. Thus, seeding of individually dispersed cells are recommended. 8. For plating, the amount of T84 cells is very crucial and, thus, should be optimal. Of note, 5  105 cells/Transwell insert (1.12 cm2) is highly recommended for growth T84 cell monolayers. Higher or lower numbers of cells would not be optimal for a monolayer establishment and for this assay. For example, T84 cell monolayers and tight junction integrity will not be developed when plating cells at a density of either 1  106 cells/Transwell insert or 1  105 cells/Transwell insert. Another important factor is the morphology of the cells seeded. This much better for indicating cell growth state. 9. In this medium renewal step, an electronic pipette aid is recommended, since the pipetting force is adjustable, and it should be set to the slowest pipetting speed. In case of using a regular pipette, gently add fresh medium into the apical chamber as slow as possible in order to avoid damaging of the monolayers. 10. Before performing cell culture medium renewal, check the color of the old cell culture medium. The cell culture medium indicator will indicate the pH. Carefully observe whether the cell culture media are acidic or basic. For instance, if the color of the culture medium in the apical chamber is bright red, this may indicate heterogeneity of the cells but also cell culture media contamination with bacteria or mycoplasma. Make sure to annotate your observations of these monolayers, which may be useful for the interpretation of unusual results.

Establishment of Intestinal Epithelial Cell Monolayers and Their Use in. . .

285

11. For every medium renewals and TER measurements, the Transwell plate containing T84 cell monolayers should always immediately be placed on a warming plate set to 37  C in laminar flow hood because tight junction assembly is temperature-sensitive [47–50]. If there is no warming plate, try to minimize handling time of the Transwell plates at room temperature. 12. In general, it is widely accepted that TER values >1000 Ω cm2 can be used for some experiments, such as short-circuit current experiment [51–53]. However, for Ca2+-switch assay, we recommend TER values of ~2500–3500 Ω cm2 (10-day monolayers). If the TER value is >4000 Ω cm2, the responsiveness to Ca2+-free S-MEM medium may be low. On the other hand, if TER value is 1.4) (Example: Nikon APO TIRF 100 1.49, Tokyo, Japan). 10. STORM/TIRF illuminator, lasers with appropriate power, and appropriate dichroics and emission filters (Example: Nikon N-STORM illuminator; LU-NV laser launch with 405, 488, 561, and 647 nm lasers; and a 405/488/561/640 m (quad pass) polychroic mirror and emission filter). Any similar system with high-power laser excitation, ability to adjust the excitation light incidence angle, a high NA objective, and a sensitive EMCCD or sCMOS camera may be utilized. 11. Software for STORM image reconstruction such as Nikon Elements or ImageJ/Fiji with the ThunderSTORM plugin [12]. 12. Image analysis software such as Nikon Elements, ImageJ/Fiji, or MATLAB.

310

3

Reena R. Beggs et al.

Methods

3.1 Buffer Preparation

1. Prepare dSTORM imaging buffers (see Note 3). Buffer A, 10 mM Tris pH 8 and 50 mM NaCl; Buffer B, 50 mM Tris pH 8 and 10 mM NaCl; and GLOX, 14 mg glucose oxidase, 50 μl catalase (17 mg/ml), and 200 μl Buffer A. The GLOX can be vortexed to completely dissolve the glucose oxidase. Buffers A and B can be stored long term at 4  C, and GLOX can be stored at 4  C for 2 weeks.

3.2 Microscope Setup

1. Select the 100 high NA objective and set the correction collar. Carefully clean the objective using lens paper and add a drop of immersion oil. If doing one color dSTORM, the bead calibration outlined in steps 2–8 is not needed; proceed directly to step 9. 2. Create the calibration sample. Dilute the 0.1 μm microspheres 1:500 in PBS (see Note 4). Add 400 μl to one well of an Ibidi 8-well chamber (or to an identical chamber as used for samples if different). 3. Clean the bottom of the chamber containing your calibration sample with ethanol and then place it on the microscope stage. Bring the microspheres into focus. 4. Remove all obstructions between the coverslip and ceiling. Turn on the 647 nm laser and set it to the lowest possible power. The laser should be visible on the ceiling, roughly directly above the objective. Adjust the laser focus to minimize the size of the spot on the ceiling. (This is typically accomplished in the software (on our system) or with a physical micrometer on the STORM illuminator.) This ensures the excitation light is focused on the objective back focal plane and is emerging from the objective parallel. 5. Use the x and y manipulators to center the laser beam so that it hits the ceiling directly above the objective. (This is typically accomplished in the software (on our system) or with a physical micrometer on the illumination arm.) This ensures the beam is aligned straight along the optical axis. Go back to step 4 to refine the focus if needed. 6. Check the focus and position of the second laser wavelength (488 nm). The alignment should be optimized for all wavelengths; repeat steps 4 and 5 if needed. 7. Find a region where there are at least 20 beads evenly dispersed in the field of view. Avoid regions with large clumps of beads or no beads. Select the calibration mode. In the imaging parameters, set the channels of interest (647 and 488), sequential image acquisition, and exposure time to 1 frame. Click run now.

dSTORM Imaging of Desmosomes

311

8. Once the calibration z-stack has been acquired, load the image in the N-STORM analysis package, click run STORM analysis, and then click “XY warp calibrate” in the identification settings menu. Close the analysis and exit the calibration mode. If you are not happy with the calibration or if it failed, go back to step 7, find a new region, and try again. Remove the calibration sample from the stage. 9. Clean and apply a drop of oil to the objective. Clean the bottom of the experimental sample with ethanol and place it on the stage. Use epifluorescence to focus on the sample. Adjust the laser alignment as outlined in steps 4 and 5. Do not rely on your previous settings as there is a difference in zoom between the calibration and image acquisition modes making realignment necessary. 3.3

Data Collection

1. Make the dSTORM buffer by combining 7 μl GLOX, 7 μl 2-mercaptoethanol, and 690 μl Buffer B on ice. Use only supernatant from the GLOX. Mix thoroughly. The dSTORM buffer should be made fresh, just before it is added to the sample (see Note 5). 2. Remove the sample buffer and replace it with dSTORM buffer. If using an 8-well Ibidi chamber, fill the well with dSTORM buffer, and place a 25 mm glass coverslip on top. This will create a closed system and prolong the life of the oxygen scavenger. 3. Find a region of interest for imaging. Look for a cell border that has multiple junctions lined up and in focus. 4. Empirically find the best illumination settings for imaging by adjusting the incidence angle of the laser (using software or a micrometer on the illumination arm). The region of interest should be bright with high contrast between the desmosomes and the background when optimal angle is achieved. Unlike some other structures commonly imaged by dSTORM, cellcell junctions are not typically visible by TIRF, and therefore the incidence angle is usually above the critical angle (see Note 6). 5. Acquire and save a widefield image of the region. 6. While the laser shutter is closed, set the laser power to 100%, and enter the number of frames to be collected in each channel (see Note 7). 7. Open the laser shutter to drive the fluorophores into the dark state, which typically takes 10–30 s. When the bulk of the fluorescence is gone and you see blinking, you have achieved the dark state. Immediately begin data acquisition (click “run now”). The images acquired will be of isolated blinking events and will be hard to interpret prior to reconstruction. Because

312

Reena R. Beggs et al.

imaging and reconstruction are both lengthy processes, it is recommended to take several images and preform a batch reconstruction. 3.4 Image Reconstruction

1. Open the image in the N-STORM analysis module (an alternative is the ImageJ ThunderSTORM plug-in). Go to a frame near the middle of the acquisition and determine the peak intensity of a single molecule. It is helpful to scroll backward and forward to find true blinking events that do not linger for more than 1–2 frames. Examine several events to get an idea for the peak heights. Once you have determined an average intensity, use it to set the minimal peak height in the identification settings. During reconstruction events with intensities below this threshold will not be included (see Note 8). This is done for each channel. 2. Run the STORM analysis. If there are multiple raw data images, you can run a batch reconstruction. 3. Once reconstruction is complete, you can further refine identification/display parameters to control which molecules are displayed in the filter settings. These allow the user to determine the number of blinking events that define a true data point, as well as narrow the radius of the defined events. For desmosomes we have found that the radius set at 10 nm and count set to 5 is optimal. 4. When the reconstructed image is satisfactory, take a “snapshot” to convert the image to a .ND2 file. Ensure the data will not be saturated by adjusting “Density Map Rendering Parameters” to display the entire dynamic range. To scale the snapshot to 4 nm/pixel, the image is set to 400% zoom, and the camera is set to 10. Save the image as a full bit-depth .tif file.

3.5

Analysis

Image analysis can be conducted in software such as Nikon Elements or ImageJ/Fiji. Our analysis methods define junction architecture by quantifying the distance between plaques and plaque length (Fig. 3) [7]. 1. Open the .tif file containing the reconstructed image in analysis software such as Fiji/ImageJ or Nikon Elements. 2. First, identify and note the locations of all puncta that are desmosomes or suspected desmosomes in the image. You can use border localization and/or co-localization with a second desmosomal protein as filters to identify candidate puncta. The following analysis will be conducted on each individual desmosome. 3. Zoom in on a puncta.

dSTORM Imaging of Desmosomes

313

4. Draw a line the length of the desmosome, along the plasma membrane. The line should measure from the beginning of any fluorescence to the end. Note the length of this line, which is the desmosome length. For a discussion of more complicated cases, see Note 9. 5. Draw a second line perpendicular to the first, bisecting the two plaques. The width of this line should be equal to the plaque length. Plot the intensity as a function of position along the line. This is the axial distribution of the protein. The distance between the two peaks represents the plaque-to-plaque distance. Output the intensity linescan to a spreadsheet. 6. Repeat steps 4 and 5 for every identified puncta. 7. Use the measurements to quantify the desmosome length and plaque-to-plaque distance. These features can be compared between treatments or labels to reveal details of desmosome architecture.

4

Notes 1. When selecting fluorophores for dSTORM, it is important to consider the duty and switching cycles, the background fluorescence, and—when doing multi-labeled experiments—the spectral overlap [13]. The duty cycle is the ratio of light and dark states of the fluorophore. Probes that have a high duty state are not ideal, as this will lead to overlap of the emission between frames, making localization difficult. The switching cycle is the average number of times a fluorophore can be switched between light and dark state before photobleaching. For obtaining structural information, it is critical to have a high switching cycle. Conversely, a low switching cycle is useful for quantification of molecules. It is also important to select probes that are photoconverted to a highly emissive state. Some fluorophores can weakly fluoresce when in a low dark state, leading to added background that does not contribute to useable events. When using multiple probes, it is important to choose spectrally distinct fluorophores to avoid cross-excitation. 2. Samples should not be on coverslips mounted on slides—they need to be in fresh dSTORM buffer at time of imaging. Another good option is 25 mm coverslips held in attofluor chambers (Invitrogen). 3. The buffer can greatly affect the properties of the probe. Most buffers employ an oxygen scavenging system, as reactive oxygen species will lead to photobleaching, and a reducing agent (thiol) to increase the lifetime of the dark state of the probe by acting as an electron donor.

314

Reena R. Beggs et al.

4. The beads will be drawn to the coverslip in PBS, while they will remain in suspension if diluted in water. 5. The dSTORM buffer should be changed just prior to imaging as the buffer works optimally for approximately 1 h. It is recommended that the sample be kept in PBS, until the setup is completed. Then the user can gently pipette off the PBS and replace it with fresh dSTORM buffer. During an experiment, when the buffer is depleted, there will be a noticeable decrease in blinking observed. At this time, it is necessary to exchange the imaging buffer. 6. Total internal reflection fluorescence (TIRF) microscopy is commonly used in SMLM because it greatly reduces the background fluorescence outside of the focal plane by restricting the illumination to ~100 nm at the coverslip/sample interface. Most desmosomes do not reside this close to the substrate and therefore will not be visible by TIRF. For imaging cell junctions, we instead use inclined illumination where the excitation laser is incident on the coverslip at an angle that is large, but still less than the critical angle needed for TIRF. This allows us to image the junctions and provides a higher signal to noise than EPI illumination (where the excitation light passes straight up though the sample). The best precise incidence angle cannot be predetermined and is determined by eye. 7. The optimal number of frames can vary widely between fluorophores and buffer systems. Once the fluorophore blinking slows significantly, there is no need to continue to collect data. We have found the ideal range when imaging Alexa Fluor 647 to be between 10,000 and 30,000 frames. 8. It is recommended to set the minimum peak height slightly less than the measured average intensity. This parameter will determine which events are selected for reconstruction, and a lower minimal peak height be more inclusive. Following reconstruction, parameters can be further refined to remove identifications desired. 9. Some desmosomes may have more complicated morphologies such as one plaque longer than the other or breaks in a plaque. The same rules should be followed while analyzing every desmosome and those rules noted in the methods. For example, if a break is less than 30 nm, the plaque is treated as contiguous, or length is always measured as the longest individual plaque.

dSTORM Imaging of Desmosomes

315

Acknowledgements This work was supported by funding to A.L.M. from the National Institutes of Health/National Institute of Arthritis and Musculoskeletal and Skin Diseases (NIH/NIAMS) (R01AR072697) and the National Science Foundation (NSF) CAREER (1832100). References 1. Pawley J (2006) Handbook of biological confocal microscopy, vol 236. Springer Science & Business Media 2. Gustafsson MG (2000) Surpassing the lateral resolution limit by a factor of two using structured illumination microscopy. J Microsc 198(Pt 2):82–87. https://doi.org/10.1046/j. 1365-2818.2000.00710.x 3. Hell SW, Wichmann J (1994) Breaking the diffraction resolution limit by stimulated emission: stimulated-emission-depletion fluorescence microscopy. Opt Lett 19(11):780–782. https://doi.org/10.1364/ol.19.000780 4. Rust MJ, Bates M, Zhuang X (2006) Subdiffraction-limit imaging by stochastic optical reconstruction microscopy (STORM). Nat Methods 3(10):793–795. https://doi.org/ 10.1038/nmeth929 5. Betzig E, Patterson GH, Sougrat R, Lindwasser OW, Olenych S, Bonifacino JS, Davidson MW, Lippincott-Schwartz J, Hess HF (2006) Imaging intracellular fluorescent proteins at nanometer resolution. Science 313 (5793):1642–1645. https://doi.org/10. 1126/science.1127344 6. McMillan JR, Haftek M, Akiyama M, South AP, Perrot H, McGrath JA, Eady RA, Shimizu H (2003) Alterations in desmosome size and number coincide with the loss of keratinocyte cohesion in skin with homozygous and heterozygous defects in the desmosomal protein plakophilin 1. J Invest Dermatol 121(1):96–103. https://doi.org/10.1046/j.1523-1747.2003. 12324.x 7. Stahley SN, Bartle EI, Atkinson CE, Kowalczyk AP, Mattheyses AL (2016) Molecular organization of the desmosome as revealed by direct stochastic optical reconstruction microscopy. J Cell Sci 129(15):2897–2904. https:// doi.org/10.1242/jcs.185785

8. Dempsey GT, Vaughan JC, Chen KH, Bates M, Zhuang X (2011) Evaluation of fluorophores for optimal performance in localization-based super-resolution imaging. Nat Methods 8(12):1027–1036. https://doi. org/10.1038/nmeth.1768 9. Huebinger J, Spindler J, Holl KJ, Koos B (2018) Quantification of protein mobility and associated reshuffling of cytoplasm during chemical fixation. Sci Rep 8(1):17756. https://doi.org/10.1038/s41598-01836112-w 10. Eltoum I, Fredenburgh J, Grizzle WE (2001) Advanced concepts in fixation: 1. Effects of fixation on immunohistochemistry, reversibility of fixation and recovery of proteins, nucleic acids, and other molecules from fixed and processed tissues. 2. Developmental methods of fixation. J Histotechnol 24(3):201–210. https://doi.org/10.1179/his.2001.24.3.201 11. Whelan DR, Bell TDM (2015) Image artifacts in single molecule localization microscopy: why optimization of sample preparation protocols matters. Sci Rep 5(1):7924. https://doi. org/10.1038/srep07924 12. Ovesny M, Krizek P, Borkovec J, Svindrych Z, Hagen GM (2014) ThunderSTORM: a comprehensive ImageJ plug-in for PALM and STORM data analysis and super-resolution imaging. Bioinformatics 30(16):2389–2390. https://doi.org/10.1093/bioinformatics/ btu202 13. Nahidiazar L, Agronskaia AV, Broertjes J, van den Broek B, Jalink K (2016) Optimizing imaging conditions for demanding multicolor super resolution localization microscopy. PLoS One 11(7):e0158884. https://doi.org/ 10.1371/journal.pone.0158884

INDEX A Absorption, distribution, metabolism and excretion (ADME)......................................................... 75 ACSF, see Artificial cerebrospinal fluid (ACSF) Acute respiratory distress syndrome (ARDS).............. 137 Adherens junctions (AJs) ..................................14, 16, 73, 123, 149, 151–153, 178 Agar bridges ........................................................ 217, 220, 223, 224, 228 Air pollution components ............................................ 152 Airway epithelial barrier (AEB) ........................... 150, 160 Airway epithelial cells AJCs ......................................................................... 149 barrier function ....................................................... 151 cell to cell barrier components ...................... 149, 150 intercellular junctions ............................................. 150 tight and adherens junctions .................................. 149 TJs ................................................................... 149–151 (see also Tight junctions (TJs)) Airway epithelial permeability air pollution components ....................................... 152 approaches, junction function ................................ 151 claudins .................................................................... 152 inflammatory cytokines........................................... 152 methods albumin .............................................................. 160 FITC ..........................................................158–159 LY (see Lucifer yellow (LY)) mannitol............................................................. 159 transepithelial/TEER (see Transendothelial electrical resistance (TEER)) two-path impedance spectroscopy ................... 158 mucosal inflammation............................................. 152 in vivo animal experiments ..................................... 151 AJs, see Adherens junctions (AJs) Albumin-alexa fluor conjugates assessment............................................... 94–95, 97–98 barrier-type endothelial cells .................................... 92 brain sections............................................................. 92 fluorescence intensity ................................................ 92 fluorochrome conjugates, antibodies....................... 91 Alveolar epithelium ..................................... 137, 138, 147 AMM, see Astrocytes Maturation Medium (AMM) AMP-activated protein kinase (AMPK) ....................... 275 Anesthesia ............................ 44, 112, 126, 131, 172, 265

Angiopoietin-1 (Ang-1) .................................50, 74, 166, 169–171, 189 Angubindin-1 .....................................292, 293, 295–298, 301, 302 Angulin-1....................................................................... 292 Angulin binder .................................................... 292, 295, 297–299, 301 Animal-based in vivo techniques.................................... 48 Anti-leak therapies......................................................... 166 Anti-TfR antibody......................................................... 204 Apical junctional complexes (AJCs)............................... 14 changes .................................................................... 152 role .................................................................. 149, 150 ARDS, see Acute respiratory distress syndrome (ARDS) Artificial cerebrospinal fluid (ACSF)................... 107, 108 Astrocytes in BBB........................................................................ 74 as co-cultures ............................................................. 50 factors......................................................................... 74 iPSC-derived.............................................................. 53 iPSC differentiation .................................................. 60 materials for differentiation, iPSC............................ 57 neurons and BMECs................................................. 74 NVU ....................................................................48, 50 Astrocytes Maturation Medium (AMM) .................57, 60 Automated measurement system ................................. 155 Avidin-biotin complex ....................................... 39, 42, 44

B Barrier-type endothelial cells ............................. 88, 91, 92 BBB, see Blood-brain barrier (BBB) BBB permeability biosafety hood and laminar flow ............................ 118 brain tissue diffusion coefficient, rat brain ............ 106 effective diffusion coefficient .................................. 119 generic fluorescence microscopy ............................ 106 health and disease.................................................... 105 measurement in vitro dextran permeability measurement .................. 109 endothelial barrier .....................................109–110 experimental treatments ................................... 108 microvessel solute permeability .............................. 120 quantitative coefficients .......................................... 106 solute tissue transport ............................................. 120 transport measurement in vivo...................... 107–108

Kursad Turksen (ed.), Permeability Barrier: Methods and Protocols, Methods in Molecular Biology, vol. 2367, https://doi.org/10.1007/978-1-0716-1673-4, © Springer Science+Business Media, LLC, part of Springer Nature 2021

317

PERMEABILITY BARRIER: METHODS

318 Index

AND

PROTOCOLS

BBB transport brain tissue............................................................... 106 non-invasive in vivo models.................................... 105 permeable membrane.............................................. 106 BECs, see Brain endothelial cells (BECs) BFGF aliquots ................................................................. 55 Bicarbonate.......................................... 217–219, 229–231 Bicinchoninic acid (BCA) ........................... 145, 294, 296 Biotin .................................................................... 207–213 Blood-brain barrier (BBB)............................................ 291 adherens junctions .................................................... 73 anatomical structure................................................ 105 astrocytes ................................................................... 74 BMECs ................................................................48, 73 brain endothelial cells ............................................... 37 brain parenchyma ...................................................... 88 cellular and molecular mechanisms ........................ 124 cerebral microvessels ............................................... 105 cerebral regions ......................................................... 37 CNS .................................................................. 47, 123 CNS diseases.............................................................. 88 CNS-targeted drug development............................. 74 co-culture model ....................................................... 51 drug development ..................................................... 74 dysfunction .............................................................. 123 endogenous transport systems ................................. 74 endothelial cells ................................................ 88, 123 endothelial layer with pericytes .............................. 105 fluorescence imaging............................................... 124 molecules ................................................................... 74 neurotherapeutics, drug development ..................... 47 pathological conditions............................................. 74 pericytes ..................................................................... 37 permeability markers ................................................................ 51 measurement ....................................................... 51 primary cells............................................................... 48 tight junctions .....................................................73, 88 transcellular pathway................................................. 88 treatment, CNS diseases ........................................... 74 in vivo models ........................................................... 48 Blood-spinal fluid barrier................................................ 74 Blood–testis barrier (BTB) blood–tissue barrier ................................................ 207 BTB integrity assay ................................208, 210–212 fluorescent images ................................................... 210 integrity in vivo ....................................................... 208 seminiferous epithelium.......................................... 207 Sertoli cell BTB integrity ............................... 208–213 in Sprague–Dawley rats........................................... 208 Blood–tissue barrier .................................... 177, 207, 212 BMECs, see Brain microvascular endothelial cells (BMECs) Bovine serum albumin (BSA)..........................38, 44, 294

Brain endothelial cells (BECs).......................37, 106, 107 angulin-1 ................................................................. 292 human stem cell-derived........................................... 50 and pericytes .............................................................. 37 porcine ................................................... 197, 198, 203 receptor antibody trafficking (see Receptor antibody trafficking in BECs) receptor-mediated endocytosis and trafficking...... 194 trafficking, endocytosed cargo ............................... 195 in vitro model, BBB ................................................ 107 Brain microcirculation ......................................... 124, 132 Brain microvascular endothelial cells (BMECs) ............ 47 angiogenesis regulation ............................................ 74 BBB......................................................................48, 74 cell sources................................................................. 49 characteristics............................................................. 74 growth factors ........................................................... 74 pericytes ..................................................................... 74 primary gatekeepers .................................................. 48 substance transport ................................................... 74 Brain microvascular pericytes ............................ 73, 75, 76 Brain microvessel isolation antibody dilution....................................................... 45 avidin-biotin complex solution ................................ 44 brain regions.............................................................. 40 BSA ......................................................................38, 44 cerebral cortex ........................................................... 44 cortical and subcortical tissue................................... 38 debris ......................................................................... 44 ethanol ....................................................................... 45 immunocytofluorescence protein immunoreactivity ............................. 42–43 solutions .............................................................. 39 optical microscope .................................................... 45 osmium tetroxide ...................................................... 44 PBS solution .............................................................. 38 pericytes ..................................................................... 38 scanning electron microscopy morphological analysis, pericyte attachment ............................................... 43–44 sucrose buffer ............................................................ 38 Western blot protein expression ......................................... 41–42 solutions ........................................................ 38–39 BSA, see Bovine serum albumin (BSA)

C Caco-2 barrier integrity .................................................. 27 Caco-2/TC7 cell line, paracellular permeability cell culture .................................................... 17, 18, 22 cell seeding ................................................................ 22 DMEM ...................................................................... 21 EGTA......................................................................... 22 FD4............................................................................ 23

PERMEABILITY BARRIER: METHODS human colon carcinoma ........................................... 15 intestinal barrier function ......................................... 17 jejunum enterocytes, lipid absorption ..................... 15 macromolecules............................................ 18, 20–21 microplate size membranes ...................................... 21 monolayer plastic support ..................................................... 15 polarized cells ...................................................... 15 null filters ................................................................... 22 pore size membranes................................................. 21 sucrase-isomaltase activity......................................... 17 TEER ...................................................... 18–19, 21–23 transcellular pathway................................................. 21 Carbachol .................................................... 218, 229, 230 Cell junctions ....................................................... 305, 311 Cell-loaded microfluidic chip ......................................... 66 Cell monolayer Caco-2 ....................................................................... 16 FD4 accumulation .................................................... 23 T84 ................................................................. 278–286 TEER ......................................................................... 18 TER................................................................. 274, 276 Central nervous system (CNS)............................... 74, 88, 105, 123, 291 Cerebral microvessels .................................................... 105 CNS, see Central nervous system (CNS) CNS disorders ........................................................ 47, 123 Co-culture model........................... 50, 51, 54, 60, 64, 67 Co-localization .................................................... 195–197, 199–205, 312 Constant phase element (CPE).................................... 300 Coomassie Brilliant Blue (CBB) ......................... 294, 296 Crumbs complex ........................................................... 151 Cytokines ................................................22, 28, 149, 152, 166, 178, 184, 259

D Data interpretation apical cell surface ..................................................... 257 creatinine flux .......................................................... 257 C. rodentium infection............................................ 259 dextran sulfate sodium (DSS)................................. 259 flux data ................................................................... 257 flux of creatinine...................................................... 261 4 kDa dextran................................................. 259, 261 kidneys ..................................................................... 261 leak pathway ............................................................ 260 mononuclear phagocyte system ............................. 261 surface area .............................................................. 257 transport .................................................................. 257 in vivo multiplex analysis ........................................ 261 Desmosomes cell junctions............................................................ 305 dSTORM ........................................................ 306–308

AND

PROTOCOLS Index 319

electron donor......................................................... 313 fluorescence microscopy ......................................... 305 fluorophore.............................................................. 313 materials.......................................................... 308–309 methods analysis .......................................................312–313 buffer preparation ............................................. 310 data collection ...........................................311–312 image reconstruction ........................................ 312 microscope setup.......................................310–311 optical microscope .................................................. 305 proteins .................................................................... 305 resolution................................................................. 305 single-molecule localizations .................................. 306 super-resolution microscopy techniques................ 305 TIRF ........................................................................ 314 Dextran permeability measurement ............................. 109 3,3’-Diaminobenzidine (DAB) ......................... 92, 98, 99 Dietary/bacterial-derived molecules................................ 2 Digital image correlation (DIC) .................................. 240 Direct stochastic optical reconstruction microscopy (dSTORM) ......................................... 306–308 Drug delivery ......................................166, 196, 291–301 receptor-mediated drug delivery to brain (see Receptor antibody trafficking in BECs) Drug permeability test, in vitro BBB models BBB modeling cell seeding ....................................................79, 80 characterization .............................................79, 80 cell culture .................................................... 76, 78–79 characterization, BBB model IF staining............................................................ 81 LY permeability ............................................. 80–81 TEER measurement......................................79, 83 drug molecule .....................................................81, 82 human cells ................................................................ 82 permeability values ..............................................82, 83 primary antibodies .................................................... 83 Ringer’s HEPES solution, drug molecule.........81, 82 secondary antibodies...........................................82, 83 triple co-culture growth medium....................... 75–76 Dulbecco’s Modified Eagle Medium (DMEM)............. 8, 10, 17, 21, 57, 75, 78, 196, 294, 301

E E-cadherin .............................................................. 16, 238 and β-catenin ........................................................... 152 in Caco2 cells .......................................................... 236 calcium-dependent adhesion .................................. 151 junctional vs. cytoplasmic localization ................... 245 localization............................................................... 244 Electrical cell-substrate impedance sensing (ECIS) airway epithelial permeability, measurement....................................... 157–158

PERMEABILITY BARRIER: METHODS

320 Index

AND

PROTOCOLS

Electrical cell-substrate impedance sensing (ECIS) (cont.) biosensor technology .............................................. 180 for EC-barrier impedance assay..................... 187–188 multifrequency provision ........................................ 181 in vitro permeability assays ....................180, 182–183 ELISA/immunoblot............................................ 137–139 Endocytosis extracellular components ........................................ 193 from luminal plasma membrane.................... 196–198 micropinocytosis ..................................................... 193 phagocytosis ............................................................ 193 pinocytosis ............................................................... 193 and subcellular receptor trafficking........................ 194 “Endohm series” ........................................................... 157 Endosomal trafficking system....................................... 193 Endothelial cells (ECs) ................................................... 50 barrier-type ................................................... 88, 91, 92 BECs (see Brain endothelial cells (BECs)) BMECs (see Brain microvascular endothelial cells (BMECs)) non-CNS ................................................................... 50 in vitro permeability cell culture .................................................181–184 ECIS assay .................................................182–183 Transwell® assay ................................................ 182 Endothelial permeability ECIS assay ..................................... 180–181, 187–188 pathophysiologic conditions................................... 178 regulator of vascular permeability ................. 178–179 routes of transportation .......................................... 178 Transwell® in vitro permeability assay barrier characteristics ........................................ 179 coating .......................................................184–185 ECs culture ................................................183–184 limitations .......................................................... 180 protocol .....................................................185–187 seeding ECs ....................................................... 185 vascular permeability ...................................... 177–179 VE-cadherin............................................................. 178 VEGF ....................................................................... 178 Endothelium............................... 47, 67, 88, 93, 177–179 Epithelial barrier maturation .......................................... 27 Epithelial cells (ECs)............................................ 274, 276 apical and basolateral membrane............................ 215 ion transport............................................................ 215 Epithelial Volt-Ohm Meter (EVOM) .................... 21, 57, 154, 276, 277, 280 Epithelium ..................................................................... 291 barrier function ....................................................... 151 intestinal ......................................................... 2, 13, 15 iPSCs.................................................................... 28–33 Sertoli cell ....................................................... 210, 211 stratified squamous ................................................. 216 Esophageal epithelial barrier basal cells ................................................................. 216

cellular pathways for ion transport......................... 216 esophageal clearance mechanisms .......................... 217 hematoxylin and eosin staining .............................. 216 pre-epithelial buffering ........................................... 217 Ussing chamber method acid treatment.................................................... 225 bicarbonate secretion measurement in pig esophagus ............................................229–230 chambers set-up ........................................220–221 circuit analysis............................................223–225 composition of stock solutions......................... 231 ion transport...................................................... 215 KCl agar bridges................................................ 220 materials.....................................................217–218 modified Ussing chamber.........................226–228 permeability measurements using fluorescein.................................................... 225 solutions ....................................................219–220 tissue dissection and mounting ................221–223 transepithelial resistance .......................... 228, 230 Evans blue dye..............................................124, 137–139 advantage ................................................................... 90 assessment............................................................ 94–96 BBB......................................................................90, 91 brain tissue................................................................. 90 fluorescence ............................................................... 90 Miles assay ...................................................... 167–170 molecular weight....................................................... 90 quantitative measurement......................................... 89 vascular permeability marker .................................... 89 EVOM, see Epithelial Volt-Ohm Meter (EVOM) Extracellular matrix (ECM)..............................54, 59, 63, 235, 236, 245

F Fetal bovine serum (FBS) ....................................... 17, 75, 161, 196, 276, 294 FITC-dextran 4 kDa (FD4) ....................................4, 6–8, 10, 18, 20, 21 Fluorescein isothiocyanate (FITC)-dextran .......... 9, 109, 158–159, 168–173 Fluorescence microscopy ................................29, 66, 106, 108, 123–133, 305, 309 Fluorescence recovery after photobleaching (FRAP) analysis.......................................................... 274 Fluorochromes ............................................................9, 92 Fluorophores .............................................. 194, 195, 197, 199, 205, 305, 306, 308, 311, 313, 314

G Glutathione S-transferase (GST).................................. 293 G-protein coupled receptor 39 (GPR39).................... 275 Graft-versus-host disease (GVHD)............................... 274 Green fluorescent protein (GFP) ........................ 294, 302

PERMEABILITY BARRIER: METHODS H High-throughput methods ............................................ 48 Horseradish peroxidase assessment..................................................... 95, 98–99 BBB permeability assessment .............................92, 93 brain capillary endothelial cells................................. 93 cerebral cortex ........................................................... 93 enzymatic activity ...................................................... 99 ultrastructural observation ....................................... 93 vascular permeability ................................................. 92 Vibratome sections.................................................... 93 hPSCs, see Human pluripotent stem cells (hPSCs) Human BMEC (hBMECs) ............................................ 68 Human colon cancer Caco-2 cells ................................. 27 Human induced pluripotent stem cells (iPSCs)............ 28 astrocytes ................................................................... 53 BMECs ...................................................................... 53 human BBB model.................................................... 48 TEER ......................................................................... 68 Human intestine .......................................................13–23 Human pluripotent stem cells (hPSCs) ...................48, 50

I ImageJ software...............................................67, 91, 113, 130, 132, 196, 197, 199–202, 242 Immunocytochemistry (ICC) ............................. 194, 196 Immunocytofluorescence .................................. 39, 42–43 Immunofluorescence (IF) staining ..............81, 196–199, 243–145, 275, 278, 282 Inflammatory bowel diseases (IBD) ............................ 274 Inflammatory mediators ........................................ 37, 166 Intestinal barrier ...........................................................1, 9, 27, 28, 273 Intestinal epithelial cell monolayers bacteria/mycoplasma .............................................. 284 Ca2+-switch assay............................................ 275, 286 cell culture medium ................................................ 284 cell types .................................................................. 274 electronic pipette..................................................... 284 FITC-dextran (4 kDa) ............................................ 286 high-resolution analyses.......................................... 274 IBD .......................................................................... 274 innate immune system ............................................ 273 lipopolysaccharides (LPS) ....................................... 274 materials Ca2+-switch assay............................................... 275 cell culture ......................................................... 278 cell culture medium and buffers....................... 275 equipment.......................................................... 277 reagents......................................................276–277 TER measurement ................................... 275, 276 methods Ca2+-switch assay.......................................280–283

AND

PROTOCOLS Index 321

protocol ............................................................. 278 T84 cell monolayers..................................278–280 peri-junctional actomyosin ring (PAMR) .............. 273 pH ............................................................................ 283 S-MEM medium ............................................ 285–286 T84 cells .................................................................. 284 TER measurements ................................................. 285 tight junction proteins ............................................ 274 tissue barrier ............................................................ 273 ultrapure/deionized water ..................................... 282 ultrastructural morphology .................................... 274 Intestinal epithelium ............................................ 2, 13, 27 Intestinal hyperpermeability ............................................. 2 Intestinal microbiota......................................................... 1 Intestinal organoids, human iPS cells antibodies, immunocytochemistry ........................... 30 cytokines and chemicals ............................................ 28 definitive endoderm ............................................29, 30 free-floating spheroids .............................................. 30 intestinal tissues......................................................... 28 materials............................................................... 28–29 monolayer culture ............................................... 30–31 monolayer epithelial formation ................................ 33 permeability assay ...................................................... 33 primers, real-time PCR ............................................. 30 TEER measurement ................................................. 31–32 value ..................................................................... 32 tight junction formation assessment ........................ 33 transepithelial barrier function assays ...................... 32 transwell insert membranes ...................................... 33 Intravital fluorescence microscopy ...................... 106, 110 In vitro BBB models aliquots preparation ............................................ 54–56 cell sources................................................................. 53 iPSC astrocytes ................................................ 57, 60, 67 BBB organ-on-chips ..................................... 63–67 hBMECs ................................................. 54, 58–59 medium preparation.................................................. 56 paracellular markers.............................................51, 52 permeability measurement........................... 57, 61–62 sucrose .................................................................51, 52 TEER measurement...............................51, 57, 60–61 transwell insert ....................................................50, 51 In vitro permeability ECs cell culture .................................................181–184 ECIS assay .................................................182–183 Transwell® assay ................................................ 182 VP assay (see Vascular permeability (VP)) In vivo BBB imaging..................................................... 124 In vivo measurement, BBB permeability animal preparation ......................................... 112–113

PERMEABILITY BARRIER: METHODS

322 Index

AND

PROTOCOLS

In vivo measurement (cont.) BECs ........................................................................ 107 concentration vs. fluorescence intensity ........ 110–111 free dye ........................................................... 117–118 generic fluorescence microscopy ............................ 106 image acquisition .................................................... 113 image analysis ................................................. 113–114 image quality ........................................................... 119 imaging system, fluorescence intensity ......... 111–112 RBCs ............................................................... 117–118 solvent drag .................................................... 117–118 2-D images ..................................................... 114–115 3-D images ..................................................... 115–117 Ion transport ........................................................ 215–217 iPSCs, see Human induced pluripotent stem cells (iPSCs) Irreversible thermodynamics ........................................ 106 Isopropyl-β-D-thiogalactopyranoside (IPTG) ............ 293

K Kedem-Katchalsky equations........................................ 106 KnockOut serum aliquots .............................................. 55

L Leak pathway................................................ 8, 14, 15, 22, 250–252, 259–261 Lipolysis-stimulated lipoprotein receptor (LSR)......... 292 Lipopolysaccharides (LPS) ........................................... 274 Lucifer yellow (LY) ........................................... 63–65, 76, 80–82, 160 Lung air/liquid barrier ................................................. 137 Lung barrier function, Evans blue dye airspaces ................................................................... 146 albumin ........................................................... 137, 146 alcohol consumption .............................................. 147 BAL fluid collection ....................................... 143, 144 fluid recovery ........................................................... 145 inflammation ........................................................... 137 lipopolysaccharide ................................................... 147 lung tissue collection ..................................... 143–144 measurement and analysis....................................... 144 mice and murine lungs ........................................... 142 permeability values .................................................. 145 serum collection .....................................140–142, 145 tail vein injection ...........................140, 141, 144, 145 tissue analysis ........................................................... 140 tissue harvest .................................................. 139, 145

M Macromolecules ................................................. 18, 20–21 Madin-Darby Canine Kidney (MDCK)....................... 274 Mannitol ....................................................................4, 159 MARVEL protein family members .............................. 151 Matrigel aliquots .......................................................55–56

Measurement in vivo............................................ 106, 107 Microelectrodes.......................................... 217, 218, 224, 226–228, 232 Microfluidic chips biological barriers...................................................... 63 endothelial permeability coefficient ......................... 67 features................................................................. 63–65 fluorescent-tagged molecule..................................... 66 human cells ................................................................ 63 molecular weight agents ........................................... 66 neurovascular unit..................................................... 66 permeability .........................................................63, 66 real-time images ........................................................ 66 transwell system......................................................... 63 vascular channel geometry........................................ 67 in vitro BBB models.................................................. 63 Micropinocytosis ........................................................... 193 Miles assay ....................................................167–168, 179 Miniaturized fluorescence microscopy, in vivo BBB permeability anesthesia ................................................................. 126 animals ..................................................................... 125 brain extravasation, endogenous tracers ................ 124 brain microcirculation............................................. 132 cocaine ............................................................ 132, 133 Evans blue................................................................ 124 fluorescence recordings image analysis .................................................... 129 miniscope........................................................... 129 habituation .............................................................. 128 imaging cannula ...................................................... 131 imaging session preparation .......................... 128–129 post-surgery care and monitoring................. 128, 131 rat and implanted cannula ...................................... 129 sodium fluorescein ......................................... 124, 131 stereotaxic cannulation surgery ..................... 126–127 surgery and imaging equipment and tools ......................................... 125 reagents and supplies ................................124–125 surgery preparation ........................................ 125–126 TEER ....................................................................... 124 Modeling barrier tissues ...........................................73–82 Molecular weight tracers ................................................ 89 Multiphoton microscopy .................................... 106, 110, 111, 113, 115

N NaFl, see Sodium fluorescein (NaFl) Near-infrared fluorescence imaging (NIRF) ............... 124 Neural Differentiation Medium (NDM).................57, 60 Neural Induction Medium (NIM)...........................57, 60 Neuronal progenitor cells (NPC) .................................. 50 Neuronal stem cell (NSC) .............................................. 50 Neurons ...............................................47, 48, 50, 74, 291

PERMEABILITY BARRIER: METHODS Neurovascular unit (NVU).......................................49, 50 NIM, see Neural Induction Medium (NIM) NIRF, see Near-infrared fluorescence imaging (NIRF) Non-CNS endothelial cells............................................. 50 Noninvasive fluorescence imaging ............................... 124 Non-invasive in vivo models......................................... 105 NPC, see Neuronal progenitor cells (NPC) NSC, see Neuronal stem cell (NSC) NVU, see Neurovascular unit (NVU)

O Ohm’s law method ........................................21, 153–154 Osmium tetroxide .............................................. 44, 95, 99

P Paracellular permeability apical junctional complex ......................................... 14 Caco-2 cell line (see Caco-2/TC7 cell line, paracellular permeability) leak pathway .............................................................. 15 leaky epithelia ............................................................ 14 molecular mechanisms .............................................. 15 TEER ......................................................................... 14 tight junctions ........................................................... 14 transcellular pathway................................................. 13 Paraformaldehyde (PFA) ................................77, 81, 196, 202, 209, 210 PDS aliquots.................................................................... 55 Pericytes ...................................................... 37, 38, 42–44, 50, 73–80, 207, 291 Peri-junctional actomyosin ring (PAMR).................... 273 Permeability coefficient................................................. 118 Permeability measurement ................................ 57, 61–62 Phagocytosis .................................................................. 193 pH stat .................................................................. 218, 229 Pig esophagus......................................217, 218, 229, 230 Pinocytosis ..................................................................... 193 Polyethersulfone (PES)................................................. 294 Polyvinylidene fluoride (PVDF)...............................41, 42 Pore pathway ................................................................... 14 Post-confluent Caco-2 cells......................................15, 16 Protein expression .....................................................41–42 Protein immunoreactivity .........................................42–43 Pulmonary edema ......................................................... 137 Pulmonary microcirculation ......................................... 137 Pulse-chase technique ................................. 194, 196, 204

R RA aliquots ................................................................55, 56 Radiometer pH stat system .......................................... 218 Radiotracer technique................................................... 106 Receptor antibody trafficking in BECs endocytosed cargo .................................................. 195

AND

PROTOCOLS Index 323

IF technique ............................................................ 194 materials antibody endocytosis from luminal plasma membrane .................................................... 196 high-resolution confocal microscope ............... 197 IF staining of subcellular markers ............196–197 ImageJ and Excel software ............................... 197 methods analysis of colocalization...........................199–202 antibody endocytosis from luminal plasma membrane ............................................197–198 IF staining of subcellular markers ............198–199 imaging of sample, confocal microscope ......... 199 quantification of co-localization score ............. 202 pinocytosis ............................................................... 193 Region of interest (ROI) .............................................. 131 Respiratory viruses ........................................................ 152

S Scanning electron microscopy ........................................ 39 Sertoli cell BTB integrity Biotinylation Reaction Mixture............................. 209, 210, 213 blood–tissue barrier ................................................ 207 functional assay........................................................ 208 materials.......................................................... 208–209 methods experiment .................................................209–210 image requisition and data analysis ..........210–212 in preparation .................................................... 209 streptavidin ..................................................... 209, 213 Short-circuit current (Isc) ..................222, 229, 230, 285 Single-molecule localization microscopy (SMLM) ..... 306 Sodium dodecyl sulfate polyacryl-amide gel electrophoresis (SDS-PAGE) ...................... 296 Sodium fluorescein (NaFl) ..................................... 52, 53, 107, 124, 132 apoMon, BBB permeability ...................................... 91 assessment..................................................... 94, 96–97 blood circulation ....................................................... 91 brain parenchyma ...................................................... 91 en bloc staining........................................................... 99 enzymatic activity ...................................................... 99 molecular weight....................................................... 91 rodent brain tissue .................................................... 91 Spermatogenesis ............................................................ 208 Stem-cell derived astrocytes............................................ 50 Stochastic optical reconstruction microscopy (STORM)............................................ 309–311 See also Direct stochastic optical reconstruction microscopy (dSTORM) Stratified squamous epithelium .................................... 216 Submucosal glands (SMG) ................................. 216, 217, 229, 230

PERMEABILITY BARRIER: METHODS

324 Index

AND

PROTOCOLS

Sucrose.................................................51, 52, 90, 97, 180 Sucrose buffer (SB) ............................................ 38, 40, 41 Super-resolution................................................... 305, 306

T Testis .............................................................................. 207 See also Blood–testis barrier (BTB) 3D-printed stretcher biological processes ................................................. 236 Caco2 cells............................................................... 236 cell-cell adhesion ..................................................... 236 characteristics........................................................... 236 computer-controlled devices .................................. 236 E-cadherin .............................................................. 236, 237, 244, 245 extracellular matrix (ECM) .................................... 235 homemade apparatuses ........................................... 236 Laminin dilution ..................................................... 243 materials.......................................................... 237–238 mechanical forces .................................................... 235 mechanical stress ..................................................... 235 methods cell culture ......................................................... 240 cell stretching ............................................240–242 fixing process .............................................240–242 imaging and analysis .................................242–243 immunofluorescence staining ........................... 242 stretching apparatus ..................................238–240 strain ........................................................................ 236 Tight junction anticoagulants.......................................................... 263 bacterial products .................................................... 251 barrier function ....................................................... 249 blood samples .......................................................... 265 Claudin-2 expression .............................................. 250 creatinine assay ...................................... 251, 266, 267 enzymatic assay........................................................ 261 epithelial cells .......................................................... 249 gastrointestinal tract................................................ 249 high-capacity flux route .......................................... 250 high protein concentrations ................................... 263 hydrogen peroxide .................................................. 261 immune-mediated colitis ........................................ 251 intestinal pore pathway ........................................... 250 intestinal unrestricted pathway............................... 262 4 kDa dextran.......................................................... 252 leak pathway ............................................................ 251 materials................................................................... 252 methods blood collection ........................................254–255 calculations ........................................................ 257 data interpretation (see Data interpretation) gavage ........................................................253–254 plasma creatinine concentration ....................... 256

plasma fluorescein .....................................255–256 plasma separation ......................................254–255 probe preparation.............................................. 253 rhodamine B fluorescence ........................255–256 protocol ................................................................... 251 retro-orbital blood .................................................. 265 small proteins .......................................................... 251 transcellular transport ............................................. 249 triple antibiotic ophthalmic ointment.................... 265 variance .................................................................... 266 in vivo assessment ................................................... 251 water-gavaged mouse.............................................. 267 wavelengths and gains............................................. 266 Tight junction formation (ZO-1 staining).................... 33 Tight junctions (TJs) .................................. 2, 73, 88, 149 E-cadherin ............................................................... 151 peripheral membrane proteins................................ 151 physical barrier function ................................ 150, 151 proteins .................................................................... 151 transmembrane protein........................................... 151 Tissue analysis................................................................ 140 Tissue edema ................................................................. 137 Tissue harvest ....................................................... 139, 145 TJ-associated MARVEL protein (TAMP) ...............4, 292 TJs, see Tight junctions (TJs) Total internal reflection fluorescence (TIRF) microscopy ................................................... 314 Transcellular pathway................................................13, 21 Transcytosis ..................................................................... 88 Transendothelial electrical resistance (TEER)................ 2, 48, 49, 51, 68, 124, 292, 299–300 advantages................................................................ 153 airway epithelial permeability, measurement ECIS ..........................................................157–158 impedance spectroscopy ...........................155–156 Ohm’s law method ...................................153–154 quantitative technique ...................................... 153 Ussing chamber.........................................156–157 Transepithelial barrier function assays ........................... 32 Transepithelial electrical measurement techniques ..... 157 Transepithelial electrical resistance (TEER) ................. 14, 18–19, 21, 27, 210, 274 Transepithelial flux ....................................................2, 158 Transepithelial resistance ..................................... 228, 230 Transepithelial voltage .................................................222, 224, 228, 230 Transferrin receptor (TfR) antibody ................... 196, 197 Trans-Golgi network (TGN)............................... 193–195 Transmembrane electric resistance (TER)................... 106 Transmission electron microscopy ................................. 99 Transport measurement in vivo ................................... 107 Transwell® assay ........................................................18, 23 coating ..................................................................... 184 setup and steps ........................................................ 183

PERMEABILITY BARRIER: METHODS in vitro permeability assay....................................... 180 barrier characteristics ........................................ 179 coating .......................................................184–185 ECs culture ................................................183–184 limitations .......................................................... 180 protocol .....................................................185–187 seeding ECs ....................................................... 185 Transwell system .......................................................51, 62 Traumatic brain injury (TBI) ......................................... 90 Tricellular tight junction angulin-1 ................................................................. 292 blood-brain barrier (BBB) ...................................... 291 Clostridium perfringens, 292 C-terminal domain.................................................. 292 epithelium................................................................ 291 Glutathione Sepharose 4B ...................................... 302 homeostasis ............................................................. 291 materials bacteria culture media....................................... 293 bacterial strains .................................................. 293 cell culture medium .......................................... 294 cell lines ............................................................. 294 flow cytometric analysis ............................294–295 plasmids ............................................................. 293 protein purification materials ...................293–294 TEER ................................................................. 295 methods cell tight junctions ....................................299–301 flow cytometric analysis ............................298–299 GST-tagged angulin modulating recombinant proteins ................................................295–297 untagged angulin modulating recombinant proteins ................................................296–298 N-terminal domains ................................................ 292 proteins .................................................................... 302 Triple co-culture........................................................75–76 TRITC-dextrans................................................................ 9 Two-path impedance spectroscopy .............................. 158

U Unsteady mass transfer model............................. 115, 119 See also BBB permeability Ussing chambers, intestinal permeability measurement advantages.................................................................... 9 and airway epithelial permeability ................. 156–157 data analysis ................................................................. 8 design....................................................................... 2, 3 dextrans........................................................................ 9 experimental conditions.............................................. 9 FD4 ............................................................... 4, 6, 7, 10 fluorescence bacteria....................................................................9 dextrans...................................................................8 tracer .......................................................................3

AND

PROTOCOLS Index 325

flux measurements....................................................... 6 genetic modifications .................................................. 4 gut permeability .......................................................... 3 gut segments ............................................................... 9 ionic fluxes, frog skin .................................................. 2 ion permeability, TEER .............................................. 2 macromolecules........................................................... 2 manometer .................................................................. 8 materials................................................................... 4–5 mouse gut .................................................................... 3 mouse intestine ........................................................... 8 mouse tissue dissection and mounting .................. 5–6 mouse tissues ............................................................... 9 occludin ....................................................................... 4 setup............................................................................. 5 solutes flow .................................................................. 2 TAMP protein ............................................................. 4 tissues ........................................................................... 8

V Vascular endothelial cadherin (VE-cadherin) ..... 178, 179 Vascular endothelial growth factor (VEGF) ...... 166, 167, 169–171, 178, 179, 189 Vascular permeability (VP) applications, VP assessment........................... 166–167 FITC-dextran ................................................. 168–173 mechanism ............................................................... 166 organ growth and performance ............................. 165 pro-and anti-VP agents........................................... 166 role ........................................................................... 165 in vivo ...................................................................... 165 FITC-dextran ............................................168–173 Miles assay .................................................167–168 Vascular permeability markers, BBB permeability albumin-alexa fluor conjugates..........................91–92, 94–95, 97–99 blood circulation ....................................................... 89 brain parenchyma ...................................................... 89 CNS diseases/disorders ............................................ 88 Evans blue dye................................. 89–91, 94–96, 99 Horseradish peroxidase................... 92–93, 95, 98–99 molecular sizes........................................................... 89 molecular weight tracers........................................... 89 sodium fluorescein .......................... 91, 94, 96–97, 99 Vesiculo-vacuolar organelles (VVOs)........................... 178 Voltage clamp .............................................. 222, 223, 228

W Western blot .................................................38–39, 41–42

Z Zonula occludens (ZO) proteins ................................151, 152, 178, 179