Vascular Hyperpermeability: Methods and Protocols (Methods in Molecular Biology, 2711) 1071634283, 9781071634288

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Methods in Molecular Biology 2711

Binu Tharakan  Editor

Vascular Hyperpermeability Methods and Protocols

METHODS

IN

MOLECULAR BIOLOGY

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

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

For over 35 years, biological scientists have come to rely on the research protocols and methodologies in the critically acclaimed Methods in Molecular Biology series. The series was the first to introduce the step-by-step protocols approach that has become the standard in all biomedical protocol publishing. Each protocol is provided in readily-reproducible step-by step 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.

Vascular Hyperpermeability Methods and Protocols

Edited by

Binu Tharakan Morehouse School of Medicine, Atlanta, GA, USA

Editor Binu Tharakan Morehouse School of Medicine Atlanta, GA, USA

ISSN 1064-3745 ISSN 1940-6029 (electronic) Methods in Molecular Biology ISBN 978-1-0716-3428-8 ISBN 978-1-0716-3429-5 (eBook) https://doi.org/10.1007/978-1-0716-3429-5 © Springer Science+Business Media, LLC, part of Springer Nature 2024 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 Caption: The microcirculation integrates a continuum of cell behaviors. The cover image shows branching PECAM+ endothelial lined vessels (green) and CD11b+ macrophages (purple) in a rat mesenteric microvascular network. The presence of both cell types emphasizes the importance of cell-cell interactions and the influence of non-vascular cells on vessel function and remodeling. (Image courtesy of Dr. Walter L. Murfee). 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. Paper in this product is recyclable.

Preface In the body, normal organ function is highly dependent on the regulation of vascular integrity and permeability. The mechanisms that regulate vascular permeability may be different in different organs and is dependent on the differences in the structure and function of the specialized microvasculature at the cellular level. Alternations that occur in normal vascular permeability, resulting in vascular hyperpermeability, contributes to the pathophysiology of many diseases that affect the central nervous system and peripheral organs. Vascular hyperpermeability is a significant complication in diseases and conditions including traumatic injuries, ischemic stroke, asthma and other inflammatory airway diseases, arthritis, chronic bowel disease, cancer, infections, and many other conditions where it can lead to impairment of organ functions and significant morbidity and mortality. Most of the earlier studies in microvascular permeability were conducted in the nineteenth century and were focused mainly on physical laws and equations, considering blood, tissue interstitial, and oncotic pressure into account. With the advent of newer imaging technologies and modern cellular, molecular, and physiological approaches, a better knowledge of various mechanisms that regulate vascular hyperpermeability, particularly in vascular cell functions and blood-vessel interactions, has emerged in the recent decades. This volume entitled Vascular Hyperpermeability: Methods and Protocols is part of the well-known Methods in Molecular Biology series, a resource book series that scientists all over the world have been using for the past several years. The focus of this volume is to organize and compile important research methods and techniques utilized to study vascular permeability/hyperpermeability in a basic science laboratory setting. The chapters listed in the volume include in vitro and in vivo approaches and conventional and modern methods. I hope this volume will allow beginners with limited experience to initiate new and affordable research projects and for established investigators to initiate new strategies and collaborations in their ongoing programs. I am very much indebted to all contributing authors for their time, hard work, and support for this project. I would like to express my gratitude and deepest appreciation to Prof. John Walker, the series editor, for his valuable help in guiding, editing, and above all his patience and understanding, without which this project would not have been successful at all. My special thanks to the office and editorial staff at Springer, particularly Ms. Anna Rakovsky and Mr. Patrick Marton, for their support and help with this project. Atlanta, GA, USA

Binu Tharakan

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

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1 Determination of Solute Permeability of Microvascular Endothelial Cell Monolayers In Vitro . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1 Jerome W. Breslin and Sarah Y. Yuan 2 Evaluation of Vascular Permeability in Inflamed Vessels of the Cremaster Muscle in Live Mice . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 13 Soi Jeong, Young Ho Choe, Young Min Kim, and Young-Min Hyun 3 Lymphatic Vascular Permeability Determined from Direct Measurements of Solute Flux . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 21 Melanie Jannaway and Joshua P. Scallan 4 Evaluation of Mesenteric Microvascular Hyperpermeability Following Hemorrhagic Shock Using Intravital Microscopy. . . . . . . . . . . . . . . . . . 39 Taylor R. Williams and Ed W. Childs 5 Determination of Endothelial Barrier Resistance by Electric Cell-Substrate Impedance Sensing (ECIS) System . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 47 Hemant Giri and Madhulika Dixit 6 Time-Lapse Observation of Cell Dynamics During Angiogenesis Using the Rat Mesentery Culture Model . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 63 Arinola O. Lampejo, Nicholas A. Hodges, Maximillian Rozenblum, and Walter L. Murfee 7 Evaluation of Barrier Integrity Using a Two-Layered Microfluidic Device Mimicking the Blood-Brain Barrier . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 77 Hossam Kadry and Luca Cucullo 8 Intravital Imaging of Leukocyte-Endothelial Interaction in Hindlimb Ischemia/Reperfusion Injury by Intravital Multiphoton Microscopy. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 89 Johannes Zeller, Karlheinz Peter, and Steffen U. Eisenhardt 9 Studying Angiogenesis Using Matrigel In Vitro and In Vivo . . . . . . . . . . . . . . . . . 105 Anantha K. Kanugula, Ravi K. Adapala, Brianna D. Guarino, Neha Bhavnani, Harshitha Dudipala, Sailaja Paruchuri, and Charles K. Thodeti 10 Determination of Blood-Brain Barrier Hyperpermeability Using Intravital Microscopy . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 117 O’lisa Yaa Waithe, Chinchusha Anasooya Shaji, Ed W. Childs, and Binu Tharakan

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Imaging and Analysis of the Dynamics of Filamentous Actin Structures in Live Endothelial Cells . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Jerome W. Breslin and Zeinab Y. Motawe Isolation and Culture of Human Umbilical Vein Endothelial Cells (HUVECs). . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Shivam Chandel, Rathnakumar Kumaragurubaran, Hemant Giri, and Madhulika Dixit Microvascular Endothelial Glycocalyx Surface Layer Visualization and Quantification . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Natascha G. Alves and Jerome W. Breslin Measurement of Blood-Brain Barrier Hyperpermeability Using Evans Blue Extravasation Assay . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . O’lisa Yaa Waithe, Xu Peng, Ed W. Childs, and Binu Tharakan Assessment of Endothelial Barrier Functions in Extra Embryonic Vasculature of Chick Embryo as an Alternative Model . . . . . . . . . . . . . . . . . . . . . . . Jamila Siamwala, Akila Swaminathan, and Suvro Chatterjee Measurement of Transendothelial Electrical Resistance in Blood-Brain Barrier Endothelial Cells . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . O’lisa Yaa Waithe, Xu Peng, Ed W. Childs, and Binu Tharakan An In Vitro Bilayer Model of Human Primary Retinal Pigment Epithelial and Choroid Endothelial Cells for Permeability Studies. . . . . . . . . . . . . . . . . . . . . . Karthikka Palanisamy and Subbulakshmi Chidambaram Quantifying Adhesion of Inflammatory Cells to the Endothelium In Vitro. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Saravanakumar Muthusamy Determination of Tight Junction Integrity in Brain Endothelial Cells Based on Tight Junction Protein Expression . . . . . . . . . . . . . . . . . . . . . . . . . . Himakarnika Alluri, Chander Sekhar Peddaboina, and Binu Tharakan Evaluation of Glycolysis and Mitochondrial Function in Endothelial Cells Using the Seahorse Analyzer . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Zeinab Y. Motawe, Salma S. Abdelmaboud, and Jerome W. Breslin Evaluation of Tight Junction Integrity in Brain Endothelial Cells Using Confocal Microscopy . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Himakarnika Alluri, Chander Sekhar Peddaboina, and Binu Tharakan

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

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Contributors SALMA S. ABDELMABOUD • Department of Molecular Pharmacology and Physiology, Morsani College of Medicine, University of South Florida, Tampa, FL, USA RAVI K. ADAPALA • Department of Integrative Medical Sciences, Northeast Ohio Medical University, Rootstown, OH, USA HIMAKARNIKA ALLURI • Precision for Medicine, Redwood City, CA, USA NATASCHA G. ALVES • Department of Molecular Pharmacology and Physiology, Morsani College of Medicine, University of Southern Florida, Tampa, FL, USA NEHA BHAVNANI • Department of Integrative Medical Sciences, Northeast Ohio Medical University, Rootstown, OH, USA; School of Biomedical Sciences, Kent State University, Kent, OH, USA JEROME W. BRESLIN • Department of Molecular Pharmacology and Physiology, Morsani College of Medicine, University of South Florida, Tampa, FL, USA SHIVAM CHANDEL • Laboratory of Vascular Biology, Centre of Excellence in Molecular Medicine, Department of Biotechnology, Bhupat and Jyoti Mehta School of Biosciences, Indian Institute of Technology Madras , Chennai, India SUVRO CHATTERJEE • Department of Biotechnology, Anna University, Chennai, India; Department of Biotechnology, The University of Burdwan, Burdwan, India SUBBULAKSHMI CHIDAMBARAM • Department of Biochemistry and Molecular Biology, Pondicherry University, Puducherry, India ED W. CHILDS • Department of Surgery, Morehouse School of Medicine, Atlanta, GA, USA YOUNG HO CHOE • Department of Anatomy, Brain Korea 21 PLUS Project for Medical Science, Yonsei University College of Medicine, Seoul, Republic of Korea LUCA CUCULLO • Department of Foundational Medical Studies, Oakland University William Beaumont School of Medicine, Rochester, MI, USA MADHULIKA DIXIT • Laboratory of Vascular Biology, Centre of Excellence in Molecular Medicine, Department of Biotechnology , Bhupat and Jyoti Mehta School of Biosciences, Indian Institute of Technology Madras, Chennai, India HARSHITHA DUDIPALA • Department of Integrative Medical Sciences, Northeast Ohio Medical University, Rootstown, OH, USA STEFFEN U. EISENHARDT • Department of Plastic and Hand Surgery, Medical Center – University of Freiburg, Faculty of Medicine, University of Freiburg, Freiburg, Germany HEMANT GIRI • Oklahoma Medical Research Foundation, Oklahoma City, OK, USA BRIANNA D. GUARINO • Department of Integrative Medical Sciences, Northeast Ohio Medical University, Rootstown, OH, USA NICHOLAS A. HODGES • J. Crayton Pruitt Family Department of Biomedical Engineering, University of Florida, Gainesville, FL, USA YOUNG-MIN HYUN • Department of Anatomy, Brain Korea 21 PLUS Project for Medical Science, Yonsei University College of Medicine, Seoul, Republic of Korea MELANIE JANNAWAY • Department of Molecular Pharmacology and Physiology, Morsani College of Medicine, University of South Florida, Tampa, FL, USA SOI JEONG • Department of Anatomy, Brain Korea 21 PLUS Project for Medical Science, Yonsei University College of Medicine, Seoul, Republic of Korea

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HOSSAM KADRY • Department of Pharmaceutical Sciences, Jerry H. Hodge School of Pharmacy, Texas Tech University Health Sciences Center, Amarillo, TX, USA ANANTHA K. KANUGULA • Department of Integrative Medical Sciences, Northeast Ohio Medical University, Rootstown, OH, USA YOUNG MIN KIM • Department of Anatomy, Yonsei University College of Medicine, Seoul, Republic of Korea; Department of Medicine, Yonsei University College of Medicine, Seoul, Republic of Korea RATHNAKUMAR KUMARAGURUBARAN • Single Cell Genomics, Cincinnati Children’s Hospital Medical Centre , University of Cincinnati School of Medicine , Cincinnati, OH, USA ARINOLA O. LAMPEJO • J. Crayton Pruitt Family Department of Biomedical Engineering, University of Florida, Gainesville, FL, USA ZEINAB Y. MOTAWE • Department of Molecular Pharmacology and Physiology, Morsani College of Medicine, University of South Florida, Tampa, FL, USA WALTER L. MURFEE • J. Crayton Pruitt Family Department of Biomedical Engineering, University of Florida, Gainesville, FL, USA SARAVANAKUMAR MUTHUSAMY • Department of Physiology, Morehouse School of Medicine, Atlanta, GA, USA KARTHIKKA PALANISAMY • R.S. Mehta Jain Department of Biochemistry and Cell Biology, Vision Research Foundation, Chennai, India SAILAJA PARUCHURI • Department of Physiology and Pharmacology, University of Toledo, Toledo, OH, USA CHANDER SEKHAR PEDDABOINA • Oxford Biotherapeutics Inc., San Jose, CA, USA XU PENG • Department of Medical Physiology, Texas A&M University College of Medicine, Bryan, TX, USA KARLHEINZ PETER • Baker Heart and Diabetes Institute, Melbourne, VIC, Australia MAXIMILLIAN ROZENBLUM • J. Crayton Pruitt Family Department of Biomedical Engineering, University of Florida, Gainesville, FL, USA JOSHUA P. SCALLAN • Department of Molecular Pharmacology and Physiology, Morsani College of Medicine, University of South Florida, Tampa, FL, USA CHINCHUSHA ANASOOYA SHAJI • College of Pharmacy, The University of Texas at Austin, Austin, TX, USA JAMILA SIAMWALA • Department of Molecular Pharmacology, Physiology and Biotechnology, Providence, RI, USA; Warren Alpert Medical School of Brown University, Providence Veterans Affairs Medical Center, Providence, RI, USA AKILA SWAMINATHAN • Department of Biotechnology, Anna University, Chennai, India; Department of Biotechnology, The University of Burdwan, Burdwan, India BINU THARAKAN • Department of Surgery, Morehouse School of Medicine, Atlanta, GA, USA CHARLES K. THODETI • Department of Integrative Medical Sciences, Northeast Ohio Medical University, Rootstown, OH, USA; School of Biomedical Sciences, Kent State University, Kent, OH, USA; Department of Physiology and Pharmacology, University of Toledo, Toledo, OH, USA O’LISA YAA WAITHE • Department of Surgery, Morehouse School of Medicine, Atlanta, GA, USA TAYLOR R. WILLIAMS • Department of Surgery, Morehouse School of Medicine, Atlanta, GA, USA SARAH Y. YUAN • Department of Molecular Pharmacology and Physiology, Morsani College of Medicine, University of South Florida, Tampa, FL, USA JOHANNES ZELLER • Department of Plastic and Hand Surgery, Medical Center – University of Freiburg, Faculty of Medicine, University of Freiburg, Freiburg, Germany; Baker Heart and Diabetes Institute, Melbourne, VIC, Australia

Chapter 1 Determination of Solute Permeability of Microvascular Endothelial Cell Monolayers In Vitro Jerome W. Breslin and Sarah Y. Yuan Abstract The microvascular endothelium has a critical role in regulating the delivery of oxygen, nutrients, and water to the surrounding tissues. Under inflammatory conditions that accompany acute injury or disease, microvascular permeability becomes elevated. When microvascular hyperpermeability becomes uncontrolled or chronic, the excessive escape of plasma proteins into the surrounding tissue disrupts homeostasis and ultimately leads to organ dysfunction. Much remains to be learned about the mechanisms that control microvascular permeability. In addition to in vivo and isolated microvessel methods, the cultured endothelial cell monolayer protocol is an important tool that allows for understanding the specific, endothelial subcellular mechanisms that determine permeability of the endothelium to plasma proteins. In this chapter, two variations of the popular Transwell culture methodology to determine permeability to using fluorescently labeled tracers are presented. The strengths and weaknesses of this approach are also discussed. Key words Monolayer permeability, Transwell permeability assay, Endothelial barrier

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Introduction The exchange of solutes and fluids across the capillary and postcapillary venule endothelium is an essential function for establishing and maintaining tissue homeostasis and optimal organ function [1, 2]. During local inflammatory responses, such as to a mosquito bite on the skin, factors released at the site of injury activate nearby endothelial cells to increase microvascular permeability and facilitate transmigration of leukocytes into the local tissues, producing localized inflammation and edema that normally resolve. However, diseases such as diabetes progressively increase microvascular permeability systemically, slowly producing a pathologic inflammatory state [3, 4]. Moreover, traumatic injuries and sepsis stimulate a systemic inflammatory response (cytokine storm) that includes systemic microvascular leakage of plasma proteins into surrounding tissues. This severe level of microvascular hyperpermeability can lead to life-threatening conditions such as acute respiratory distress

Binu Tharakan (ed.), Vascular Hyperpermeability: Methods and Protocols, Methods in Molecular Biology, vol. 2711, https://doi.org/10.1007/978-1-0716-3429-5_1, © Springer Science+Business Media, LLC, part of Springer Nature 2024

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syndrome, abdominal compartment syndrome, and multiple organ dysfunction [5–11]. The passage of fluid and solutes across the microvascular wall is determined by (1) the net hydrostatic and osmotic pressure gradients across the semipermeable endothelial barrier and (2) the barrier properties of endothelium [1]. Under physiological conditions, the pressure gradient across the endothelium produces a small, net filtration of fluid from the blood into the surrounding tissues. The physiological, continuous formation of interstitial fluid serves to deliver nutrients to the cells of the tissues and is cleared by lymphatic drainage [12, 13]. When considering flux of this fluid across the microvascular wall, the capillary filtration coefficient introduced by Landis, a product of the hydraulic conductivity and surface area for exchange, is used to describe the rate of water passage through the endothelium [14]. For solutes, their transport across the endothelium can be described by a few different terms. First, the osmotic reflection coefficient is a term introduced by Kedem and Katchalsky into the Landis equation that indicates the fraction of proteins that are retained in plasma during transendothelial filtration of fluid, i.e., essentially the inverse of the fraction of plasma proteins that escape across the endothelial wall [15]. More commonly, terms that describe the diffusive transport of solutes across the endothelium are used. These include the solute flux (Js), which is the amount of solute that crosses the endothelium per unit of time, and the solute permeability coefficient (Ps), a one-dimensional rate indicating how quickly solute crosses the endothelium. By convention, the units for Ps are reported as cm/s. The relationship between Js and Ps is defined in Fick’s first law of diffusion: J s = P s S ðC PL - C ISF Þ This relationship indicates that the net Js for a particular solute across the endothelium is the product of Ps, the surface area of the endothelium available for exchange (S), and the concentration gradient of the solute across the endothelium, established by the difference of its plasma concentration (CPL) minus its concentration in the interstitial fluid (CISF). It is important to note that Fick’s first law describes the relationship for diffusive permeability properties of the endothelium in the absence of any hydrostatic or osmotic pressure gradients between the plasma and interstitial fluid compartments. An increase in the endothelium’s Ps to a particular solute or group of solutes will facilitate their leakage from plasma into the surrounding tissues. However, it is worth noting that in vivo, increased appearance of plasma solutes in the surrounding tissues can also be elicited by increased filtration of plasma due to increased local capillary hydrostatic pressure and capillary recruitment, both which result from increased local blood flow. The ability to assess Ps thus represents the contribution of the endothelium’s

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diffusive permeability properties to microvascular leakage, independent of changes in pressure gradients across the endothelium [1, 2]. While there are several useful in vivo and isolated microvessel models that can be used to estimate or determine Ps, the establishment of culture methods for endothelial cells starting in the early 1970s [16] has facilitated development of a variety of protocols to study transport of solutes across the endothelium. One very popular method is to grow endothelial cells into a monolayer on semipermeable membranes and then place the membrane between two reservoirs, one that has tracer and the other on which accumulation of the tracer is measured (Fig. 1). The tracers may be labeled radioactively, are sometimes detected with chemical means, but most commonly are labeled with fluorophores. The data can then be presented simply as the amount of tracer that accumulates over a given time period. However, if the concentration of the tracer is measured on both sides of the endothelial monolayer, then Fick’s first law of diffusion can also be used to calculate the apparent Ps of the tracer. While the major drawback to endothelial monolayer models is that they do not fully represent the in vivo microcirculation or intact microvessels under constant intravascular flow and perfusion pressure, they do offer the advantage of providing information about endothelial-specific mechanisms that contribute to the control of microvascular permeability. In addition, the protocol is relatively easy to perform and allows other advantages of endothelial cell culture models such as the ability to genetically modify cells or use chemical inhibitors of cell function that might not be useful in the in vivo setting. Several considerations should be made when planning an experiment to determine the solute permeability of endothelial monolayers in vitro. While some primary endothelial cells are very easy to grow, like human umbilical vein endothelial cells (HUVEC), microvascular endothelial cells may better represent the microcirculation. In addition, endothelial cells chosen from a particular organ might be more representative of that organ’s microvascular beds and are available for a variety of organs, including the lung, heart, skin, brain, intestine, and others [5, 17– 26]. The culture conditions are a second important consideration. Ideally, the semipermeable membrane upon the cells are grown should be coated with gelatin, fibronectin, or a mixture of basement membrane proteins that will promote good focal adhesion. In addition, the cells should be seeded at a density to immediately achieve confluence, and the monolayer should be allowed to grow for 5–7 days to allow junctions to mature [22, 24]. The choice of tracer is also important. Albumin that is labeled with a fluorophore is often chosen because albumin is one of the major plasma components. Choice of an albumin that is labeled with a 1:1 ratio of fluorophore is desirable to achieve a linear standard curve for fluorescence intensity and protein concentration. Some types of

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Fig. 1 Endothelial cell monolayer permeability cultureware and chamber systems. (a) Image of a 6-well culture plate with Costar Snapwell membranes on which the cells are grown. The Costar Transwell membranes (not shown) described in this protocol are similar, but smaller, in a 24-well plate configuration. (b) Schematic showing the configuration of the upper, “luminal” and lower, “abluminal” compartments with the endothelial cells grown on the Transwell insert in between. A tracer is added to the top compartment, and the rate of appearance of tracer in the lower compartment is measured to determine permeability of the cell monolayer.

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FITC-albumin that are commercially available have multiple FITC molecules attached, some by noncovalent binding, to make the fluorescence brighter per molecule. However, these present the problem that some FITC molecules can dissociate from the albumin, cross the endothelial barrier, and make the apparent Ps much higher than the actual Ps. Another consideration is that certain fluorophores can alter the physiochemical characteristics of tracer molecules such as albumin [27]. FITC was shown to significantly alter the size and relative charge of albumin, and FITC-albumin flux was recorded to be significantly higher than TRITC-flux in single-perfused microvessels [27]. Some of these problems may be alleviated with the newer Alexa Fluor fluorophores, which are brighter and represent a better choice for obtaining 1:1 labeling with albumin. In addition, dextrans are labeled 1:1 with fluorophore and can be obtained in various molecular weight ranges, which allow for determining the permeability of molecules of different sizes [21, 28]. Lastly, consideration should be given to whether a protocol that provides a single Ps per experimental group or one that compares Ps of the same endothelial monolayers before and after an experimental intervention is needed to satisfy the study objectives. An advantage of the variation in which a single Ps is determined is that the cultureware used requires a relatively small amount of media and tracer because the cell surface-coated membranes are small (Fig. 1b). In contrast, the variation in which multiple measurements of solute flux over time can yield values of Ps before and after addition of test compounds utilizes a specialized chamber system that has the drawback that it requires larger amounts of media and tracer (Fig. 1c). However, this latter method has the advantage that it can provide detailed data about changes in Ps over time. Both variations of the protocol are discussed in this chapter.

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Materials 1. Microvascular endothelial cells and their recommended growth medium. 2. Phenol red-free endothelial basal medium (e.g., EBM-PRF from Lonza or VBM from Lifeline Cell Technology). 3. Phosphate-buffered saline (PBS). 4. 0.25% trypsin-EDTA solution.

ä Fig. 1 (continued) (c) Schematic of the configuration of the Ussing Snapwell chamber system, in which the “luminal” compartment is on the left and the “abluminal” compartment is on the right. Transport of the tracer from left to right across the endothelial monolayer is measured in order to determine permeability

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5. Hemacytometer. 6. Gelatin solution (1.5% in 0.9% NaCl) or another matrix solution of your choice. 7. Costar Transwell Membranes (0.4 μm pores, clear polyester membrane): (a) For single Ps determination: Corning #3470, twelve 0.33 cm2 inserts in a 24-well plate. (b) For multiple Ps determination: Corning #3801, six 1.13 cm2 inserts in a 6-well plate. 8. For multiple Ps determination, a special chamber system with heating manifold base assembly and a circulating water bath are required (NaviCyte Ussing vertical chamber system available from Warner Instruments; part number 66-0075 for the base assembly; part number 66-0008 for Snapwell chambers). 9. Tracer of your choice. Some examples are FITC-Albumin, Alexa Fluor-488-Albumin, FITC-Dextran-70-kDa, TRITCDextran-4.4-kDa, and TRITC-Dextran-150-kDa. 10. Black, solid 96-well plate, and a plate reader to measure fluorescence, such as a Molecular Devices SpectraMax L Microplate Reader. 11. Distilled or Millipore-filtered water for diluting samples in the 96-well plate.

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Methods

3.1 Preparation of Endothelial Cell Monolayers for Study

1. Grow endothelial cells to 80–90% confluency in 60-mm or 100-mm plates in their recommended growth medium. 2. Warm growth medium, 0.25% trypsin-EDTA solution, PBS, and gelatin solution to 37 °C. 3. In a biosafety cabinet, coat the Transwell or Snapwell membrane inserts with gelatin solution. For 0.33 cm2 Transwell inserts, apply 100 μL per well. For 1.13 cm2 Snapwell inserts, add 500 μL per well. After applying to all of the wells, the solution may be aspirated, leaving a thin coat on the membranes. 4. Cover the plate containing the inserts, set aside, and bring the plate of cells to be trypsinized to the biosafety cabinet. 5. Aspirate the growth medium and add 2 mL of warm PBS. 6. Aspirate the PBS and add 1–2 mL of warm 0.25% TrypsinETDA solution. 7. Allow the cells to round up, and rap the dish on the bench if necessary to get a cell suspension.

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8. Add 2 mL of warmed growth medium to the cells to inactivate the trypsin. Collect the cell suspension from the plate with a pipette and place into a 15-mL conical tube. 9. Wash the dish with 2 mL of PBS to collect any remain cells, and then pipette this suspension into the same 15-mL conical tube. 10. Take a 10 μL of the cell suspension, and place in the hemacytometer. Perform a cell count while waiting in the next step. 11. Centrifuge the cells at 300×g for 3 min. 12. Resuspend the cells in an appropriate amount of growth medium, taking into account that the seeding density for the membranes is approximately 105 cells per cm2 surface area (see Note 1). Seed the following: (a) For each Transwell insert (0.33 cm2), seed 3 × 104 cells in 100 μL of medium. (b) For each Snapwell insert (1.13 cm2), seed 105 cells in 500 μL of medium. 13. Add growth medium to the bottom wells supporting each insert: (a) For the wells containing Transwell inserts, add 600 μL of medium. (b) For the wells containing Snapwell inserts, and 2 mL of medium. 14. Place in the incubator, and allow the cells to grow for 5–7 days, performing a medium change within 24 h of seeding and then every 48 h thereafter (see Note 2). 3.2 Assay Variation 1: Transwell Membranes

1. Warm the phenol red-free basal medium to 37 °C. 2. In the biosafety cabinet, aspirate the growth medium from both the inserts (upper compartment) and the wells under them (lower compartment). 3. Replace the medium with the phenol red-free basal medium (100 μL in the upper compartment and 600 μL in the lower compartment), and place the cells in the incubator. 4. Allow the cells at least 1 h to equilibrate in the new media before starting the experiment. 5. Prepare 200 μL of 10 μg/mL FITC-albumin (or other tracers of your choice) in phenol red-free medium. 6. Depending upon the protocol, add the inhibitors or agonists that are hypothesized to increase or decrease permeability at the chosen time points before the tracer will be added (see Note 3).

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7. Add 10 μL of the tracer stock solution to the each of the upper compartments. Place the plate back in the incubator and wait 30 min (see Note 4). 8. Set up the standard curve dilutions in the 96-well black plate while the cells are incubating. In column 1, row A, add 196 μL of distilled water, and then to that add 4 μL of the 10 μg/mL FITC-albumin solution. In rows B through H, add 100 μL. Then remove perform serial dilutions starting with removal of 100 μL from well A1 and transferring it to well B1, mixing, then removing 100 μL from well B1 and adding to well C1, etc. until reaching well G1. Discard the 100 μL removed from well G1. Leave well H1 as a blank. Keep the plate covered until samples are added later. 9. After 30 min, get the plate from the incubator, and move the Transwell inserts to the empty wells to stop flux of tracer from the upper compartment to the lower compartment. 10. Take 20 μL samples in duplicate from each of the lower compartments, and load them into the black, solid 96-well plate. Add 80 μL of water to each well containing a sample. From the upper compartment, take 10 μL samples in duplicate, place them in the plate, and mix with 90 μL of water. 11. Determine the concentrations of the upper compartment (luminal concentration) and the lower compartment (abluminal concentration) using the plate reader and standard curve, taking dilutions into consideration. 12. Calculate Ps for each well using the following equation: P s = ðC A =t Þ × ð1=AÞ × ðV=C L Þ where: CA = albumin concentration (mg/mL) t = time in seconds A = area = 0.33 cm2 V = volume of the abluminal chamber (0.6 mL; see Note 5) CL = luminal concentration (mg/mL). 13. This Ps value represents the average Ps for the time period between the addition of tracer to the luminal compartment and the end of the experiment when the abluminal samples were taken. 3.3 Assay Variation 2: Snapwell Membranes

1. Warm 100 mL of phenol red-free basal medium to 37 °C. 2. Turn on the water bath that warms the Ussing chamber base assembly.

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Fig. 2 Snapwell chamber assembly. (a) Attachment of the retaining ring to lock both sides of the chambers together. The yellow arrow shows the retaining ring, held by the attachment tool. A ring must be placed on both sides. Note that both compartment openings are on top. (b) Attachment of the tubes for CO2 delivery to the chambers. The yellow arrows show attachment points, and adjustment knobs are on the other end of the tubing on the manifold

3. In the biosafety cabinet, one at a time, aspirate the growth medium from an insert, remove it from the plate, and then insert it between the Ussing chamber half-blocks, and place a retainer ring on each side (Fig. 2a). Immediately add 5 mL of warm media to each side, and place the chamber block into the base assembly. Repeat for all six inserts. 4. Attach the gas tubes that deliver 5% CO2 to the tops of each chamber (Fig. 2b). Open the CO2 tank, and use the adjusters on the base assembly to regulate bubbling in all of the chambers. 5. Allow the cells to equilibrate for 30 min. 6. While waiting, prepare 6 mL of a 50 mg/mL FITC-albumin in phenol red-free basal medium. 7. Set up a standard curve for FITC-albumin in solid black 96-well plate as follows: in column 1, row A, add 199.2 μL of distilled water, and then to that add 0.8 μL of the 50 μg/mL FITC-albumin solution. In rows B through H, add 100 μL. Then remove perform serial dilutions starting with removal of 100 μL from well A1 and transferring it to well B1, mixing, then removing 100 μL from well B1 and adding to well C1, etc. until reaching well G1. Discard the 100 μL removed from well G1. Leave well H1 as a blank. Keep the plate covered until more samples are added. 8. In each chamber, identify the “luminal” side on which the apical surface of the monolayers face. Remove 1 mL of medium from this side, and replace with 1 mL of the 50 mg/mL FITCalbumin solution. This will make the luminal concentration

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10 mg/mL for FITC-albumin. Take two 20 μL aliquots from each luminal compartment and place in the 96-well plate. 9. To determine baseline flux, take 20-μL samples in duplicate from each abluminal compartment every 5 min (see Notes 6 and 7) for the predetermined baseline period (usually 30 min or 60 min). Keep the 96-well plate covered when not adding samples (see Note 8). 10. Add the test compound(s) to each well, and continue collecting samples for the periods predetermined by the experimental design. 11. At the end of the experiment, take two 20-μL aliquots from each luminal compartment again. 12. Bring all the volumes of the samples taken up to 100 μL by adding 80 μL water. Determine protein concentrations using a fluorescent plate reader, taking the dilutions of samples into consideration. 13. Create a spreadsheet in which the albumin protein concentrations for each chamber are plotted over time. The slope of the line between two time points multiplied by the abluminal compartment volume represents the flux of the tracer for the period between those two time points. 14. Calculate Ps for each time point of each chamber as follows: P s = ½ðC A1 - C A0 Þ=t  × ð1=A Þ × ðV =C L Þ where: CA0 = albumin concentration at previous time point (mg/mL) CA1 = albumin concentration at current time point (mg/mL) t = time between the two time points, in seconds, A = area = 1.13 cm2 V = volume of the abluminal chamber (5 mL; see Note 5) CL = average luminal concentration (mg/mL) from measurements at the beginning and end of the experiment. 14. The values for Ps calculated this way represent the average for the time period between time points. This approach allows for determining changes in Ps over time.

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Notes 1. One well may be left empty with no cells added, only growth medium, to serve as a positive control for maximal permeability achieved through the membrane alone. 2. In some cases, such as when cells are transfected with plasmid DNA just prior to seeding, a shorter incubation period may be required due to the transient nature of some expression vectors. 3. Some compounds elicit increases in permeability within seconds and are short-lived (e.g., histamine and VEGF), while others take several hours to reach their maximal increase (e.g., IL-1β). This, plus any multistep protocols in which inhibitors are added, should be taken into consideration when designing protocols. 4. A longer or shorter time period can be used, if desired. 5. Note 1 mL = 1 cm3. 6. Time points can be farther apart (e.g., 10 min.) if desired. Note that the Ps will represent the average Ps for each time interval. 7. After collecting two 20 μL samples from the abluminal chamber, a 40 μL volume of medium of phenol red-free basal medium should be added to maintain the overall volume of the compartment, and prevent differences in volume between the luminal and abluminal compartments. 8. Additional 96-well plates may be needed if the number of samples obtained is large.

Acknowledgments The authors acknowledge their support from NIH grants NIH R35 HL1507321, R01GM097270 (SYY), and R01GM120774 (JWB). References 1. Dura´n WN, Sa´nchez FA, Breslin JW (2008) Microcirculatory exchange function. In: Tuma RF, Dura´n WN, Ley K (eds) Handbook of physiology: microcirculation. Academic Press/Elsevier, San Diego, pp 81–124 2. Yuan SY, Rigor RR (2010) Regulation of endothelial barrier function. San Rafael, Morgan & Claypool Life Sciences 3. Yuan SY, Breslin JW, Perrin R et al (2007) Microvascular permeability in diabetes and insulin resistance. Microcirculation 14:363– 373

4. Yuan SY, Ustinova EE, Wu MH et al (2000) Protein kinase C activation contributes to microvascular barrier dysfunction in the heart at early stages of diabetes. Circ Res 87:412– 417 5. Alves NG, Trujillo AN, Breslin JW et al (2019) Sphingosine-1-phosphate reduces hemorrhagic shock and resuscitation-induced microvascular leakage by protecting endothelial mitochondrial integrity. Shock 52:423–433 6. Beard RS Jr, Yang X, Meegan JE et al (2016) Palmitoyl acyltransferase DHHC21 mediates

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endothelial dysfunction in systemic inflammatory response syndrome. Nat Commun 7: 12823 7. Breslin JW, Daines DA, Doggett TM et al (2016) Rnd3 as a novel target to ameliorate microvascular leakage. J Am Heart Assoc 5: e003336 8. Breslin JW, Wu MH, Guo M et al (2008) Tolllike receptor 4 contributes to microvascular inflammation and barrier dysfunction in thermal injury. Shock 29:349–355 9. Doggett TM, Alves NG, Yuan SY et al (2017) Sphingosine-1-phosphate treatment can ameliorate microvascular leakage caused by combined alcohol intoxication and hemorrhagic shock. Sci Rep 7:4078 10. Huang Q, Xu W, Ustinova E et al (2003) Myosin light chain kinase-dependent microvascular hyperpermeability in thermal injury. Shock 20: 363–368 11. Reynoso R, Perrin RM, Breslin JW et al (2007) A role for long chain myosin light chain kinase (MLCK-210) in microvascular hyperpermeability during severe burns. Shock 28:589–595 12. Breslin JW (2014) Mechanical forces and lymphatic transport. Microvasc Res 96:46–54 13. Breslin JW, Yang Y, Scallan JP et al (2019) Lymphatic vessel network structure and physiology. Compr Physiol 9:207–299 14. Landis EM (1927) Microinjection studies of capillary permeability. II. The relation between capillary pressure and the rate at which fluid passes through the walls of single capillaries. Am J Phys 82:217–238 15. Kedem O, Katchalsky A (1958) Thermodynamic analysis of the permeability of biological membranes to non-electrolytes. Biochim Biophys Acta 27:229–246 16. Jaffe EA, Nachman RL, Becker CG et al (1973) Culture of human endothelial cells derived from umbilical veins. Identification by morphologic and immunologic criteria. J Clin Invest 52:2745–2756 17. Tinsley JH, Breslin JW, Teasdale NR et al (2005) PKC-dependent, burn-induced adherens junction reorganization and barrier dysfunction in pulmonary microvascular endothelial cells. Am J Physiol Lung Cell Mol Physiol 289:L217–L223 18. Breslin J, Pappas P, Cerveira J et al (2003) VEGF increases endothelial permeability by

separate signaling pathways involving ERK-1/ 2 and nitric oxide. Am J Phys Heart Circ Phys 284:H92–H100 19. Breslin JW, Sun H, Xu W et al (2006) Involvement of ROCK-mediated endothelial tension development in neutrophil-stimulated microvascular leakage. Am J Physiol Heart Circ Physiol 290:H741–H750 20. Breslin JW, Yuan SY (2004) Involvement of RhoA and Rho kinase in neutrophil-stimulated endothelial hyperpermeability. Am J Physiol Heart Circ Physiol 286:H1057–H1062 21. Alves NG, Yuan SY, Breslin JW (2019) Sphingosine-1-phosphate protects against brain microvascular endothelial junctional protein disorganization and barrier dysfunction caused by alcohol. Microcirculation 26:e12506 22. Breslin JW, Pappas PJ, Cerveira JJ et al (2003) VEGF increases endothelial permeability by separate signaling pathways involving ERK-1/ 2 and nitric oxide. Am J Physiol Heart Circ Physiol 284:H92–H100 23. Rigor RR, Beard RS Jr, Litovka OP et al (2012) Interleukin-1beta-induced barrier dysfunction is signaled through PKC-theta in human brain microvascular endothelium. Am J Physiol Cell Physiol 302:C1513–C1522 24. Beard RS Jr, Haines RJ, Wu KY et al (2014) Non-muscle Mlck is required for beta-cateninand FoxO1-dependent downregulation of Cldn5 in IL-1beta-mediated barrier dysfunction in brain endothelial cells. J Cell Sci 127: 1840–1853 25. Beard RS Jr, Hoettels BA, Meegan JE et al (2020) AKT2 maintains brain endothelial claudin-5 expression and selective activation of IR/AKT2/FOXO1-signaling reverses barrier dysfunction. J Cereb Blood Flow Metab 40:374–391 26. Robinson BD, Shaji CA, Lomas A et al (2018) Measurement of microvascular endothelial barrier dysfunction and hyperpermeability in vitro. Methods Mol Biol 1717:237–242 27. Bingaman S, Huxley VH, Rumbaut RE (2003) Fluorescent dyes modify properties of proteins used in microvascular research. Microcirculation 10:221–231 28. Motawe ZY, Farsaei F, Abdelmaboud SS et al (2020) Sigma-1 receptor activation-induced glycolytic ATP production and endothelial barrier enhancement. Microcirculation 27:e12620

Chapter 2 Evaluation of Vascular Permeability in Inflamed Vessels of the Cremaster Muscle in Live Mice Soi Jeong, Young Ho Choe, Young Min Kim, and Young-Min Hyun Abstract Inflammation in vascular structures due to external factors such as injury or infection inevitably leads to blood leakage. Therefore, measuring blood infiltrated into tissue may serve as an indication for the extent of an inflammatory reaction or injury. There are various methods of confirming vascular permeability in vivo and in vitro; for example, using a blood vessel permeable dye, the dye efflux can be quantitatively measured with a spectrophotometer. Although the aforementioned commonly used methods can measure leaked dye without difficulty, substantial limitations exist regarding the time points of blood leakage that can be measured. Here, we describe the details of a novel protocol to identify and analyze the real-time progression of blood leakage in vivo. This method, by combining existing methods with real-time imaging, is expected to immensely improve the visualization and evaluation of vascular permeability. Key words Cremaster muscle exposure surgery, Two-photon intravital imaging, Vascular permeability

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Introduction Increased vascular permeability is a key event that accompanies inflammatory reactions [1]. Blood vessels in the inflamed area are relaxed, and the permeability of the capillaries is increased [2]. Such phenomena results in increased outflow of blood toward the site of inflammation, resulting in fluid accumulation and edema in tissues [3]. In addition, increased vascular permeability can induce immune cell migration and outflow through blood leakage, resulting in an excessive immune response [4–7]. Regulation of vascular permeability also plays an important role in maintaining body homeostasis [8]. Therefore, the study of vascular permeability has been important in terms of disease regulation and mitigation [9– 11]. Vascular permeability can be measured via blood outflow using various methods, such as hydraulic conductivity, endothelium electrical resistance, and albumin transport [12]. In addition,

Binu Tharakan (ed.), Vascular Hyperpermeability: Methods and Protocols, Methods in Molecular Biology, vol. 2711, https://doi.org/10.1007/978-1-0716-3429-5_2, © Springer Science+Business Media, LLC, part of Springer Nature 2024

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experiments utilizing human microvascular endothelial cells (hMVEC) may be conducted to mimic the microvascular environment [13]. These methods were an attempt to reconstruct in vivo experimental conditions; however, actual in vivo phenomena that occur in arbitrarily set environments are hard to detect using such methods. The most frequently used experimental method to show blood leakage in vivo utilizes Evans blue dye [14]. Evans blue dye binds to albumin when injected via the tail vein. As inflammatory reactions occur, blood vessel permeability is increased, and albumin-bound Evans blue dye diffuse from the blood vessels out to the tissue. In other words, by detecting absorbance in tissue in which Evans blue dye has spread, one can confirm that blood leakage has indeed occurred. However, vascular permeability measurement with Evans blue dye is available only at a single time point; therefore, it is not a viable option for the monitoring fluid outflow in a real-time manner. Here, we describe a method for the detection and analysis of blood leakage in murine blood vessels in real time, using intravital imaging [15]. The blood vessels of the cremaster muscle were examined by two-photon microscopy to measure blood outflow and the neutrophil migration in the blood vessels of inflamed mice. Prior to intravital imaging, mice were injected with Texas Red dextran through the tail vein. Since injected Texas Red dextran follows the bloodstream, real-time imaging on Texas Red dextran leaking out of the blood vessel may serve as evidence of blood leakage due to inflammation. Through analysis of the obtained imaging data, the time and intensity of vascular leakage can be visualized and quantified. Thus, combining other experimental methods and real-time imaging to measure vascular permeability may be considerably useful, providing information on blood vessel dysfunction during inflammatory conditions.

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Materials

2.1 Imaging Chamber Preparation

1. Prepare an empty plastic frame to establish imaging chamber (see Note 1). 2. Place a thick plastic sheet in the middle of the frame to secure room for the mouse (see Note 2). 3. Mix curing agent and base of SYLGARD 184 silicone elastomer kit (Dow Corning) in a different container at a ratio of 1: 10, and pour mixture into frame (see Note 3). 4. Harden for about a day in a 50 °C incubator to promote polymerization. 5. Carefully remove plastic sheet.

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Fig. 1 Overview of cremaster muscle imaging chamber

6. Mix SYLGARD reagents in the same ratio as step 3 and pour into a 12-well plate to create a pedestal. During imaging, the murine cremaster muscle will be placed and fixed on this pedestal (see Note 4). 7. Similarly, after hardening in a 50 °C incubator for a day, carefully lift the silicone with a pointed tool. 8. Part of the pedestal is cut with a blade to position it in the center of the premade plastic frame. 9. Fix the bottom part of pedestal to the plastic frame using super glue. 10. After about a day, the pedestal is completely attached and may be used as an imaging chamber (Fig. 1). 2.2 Blood Vessel Labeling

1. Prepare dextran (MW: 70 kDa) conjugated with Texas Red dye (Invitrogen) (see Note 5). 2. Powdered dextran is added to sterile 1× PBS to reach a final concentration of 2.5 mg/mL. 3. Filter dissolved dextran solution using a 0.22 μm syringe filter and transfer to new tube. 4. Dispense 500 μL each into a light-blocking Eppendorf tube and store at -20 °C.

2.3

Instruments

1. Select a suitable microscope for intravital imaging. In this paper, LSM 7MP (Zeiss) was used (see Note 6). 2. For long-term imaging, isoflurane is applied using a respiratory anesthetic machine (Kent Scientific). 3. For analysis of imaging data, Volocity v6.3.1 (PerkinElmer) is used, and Prism v7.0 (GraphPad) is used for graph construction.

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Method

3.1 Cremaster Muscle Exposure Surgery

1. Prior to surgery, zolazepam/tiletamine (30 mg/kg) and xylazine (10 mg/kg) are mixed and then injected into the abdominal cavity of the mouse. The injected amount is within 10% of the mouse weight. 2. Before exposing the cremaster muscle, scrotal fur is removed with hair removal cream (see Note 7). 3. Place the mouse in the center of the imaging chamber and fix the legs and tail with surgical tape. 4. Place the scrotum on the pedestal, and pull the skin with a suture needle until it is fully stretched. 5. Anchor the tight suture thread on the floor with surgical tape and drop prewarmed 1× PBS around the scrotum (Fig. 2a). 6. Using a dissecting microscope, cut the edge of the stretched skin with scissors (Fig. 2b, c). 7. Carefully remove the connective tissue inside the skin with microdissection scissors to expose the cremaster muscle (Fig. 2d). 8. Forceps are used to push the testicle out of the body to expose the cremaster muscle. 9. Fix the exposed cremaster muscle on the pedestal using an insect pin (Fig. 2e) (see Note 8).

Fig. 2 Step of cremaster muscle exposure surgery. (Reproduced from Ref. [15] with permission from Hindawi)

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10. The skin is taken out of the pedestal and then dropped with prewarmed 1× PBS to prevent the cremaster muscle from drying out. 11. Carefully make an incision along the middle of the cremaster muscle with microdissection scissors (Fig. 2f). 12. Attach cremaster muscle on the pedestal with insect pins, and tuck the exposed testicle carefully into the body using forceps (Fig. 2g). 13. Fill the imaging chamber with prewarmed 1× PBS to moisten the cremaster muscle fixed on the pedestal (Fig. 2h). 3.2 Two-Photon Intravital Microscopy

1. Prior to imaging, a premade dextran solution is injected intravenously at 25 mg/kg to visualize blood vessels (see Note 9). 2. Place the imaging chamber on a heating plate to control the body temperature of the mouse. 3. Connect the nose cone to the mouse for a constant supply of breathing anesthesia. 4. Turn on the laser of the two-photon microscope, adjust the power, and then place the lens on the cremaster muscle and perform imaging (see Note 10).

3.3 Vascular Permeability Measurement

The ZEN file obtained using a two-photon microscope is imported into Volocity software for analysis. 1. Upon starting the program, click on “Create a new library” to create a new library (see Note 11). 2. Add imaging data to the newly made library. 3. Noise can be removed from raw data by selecting “Remove noise” from the “Tools menu” (see Note 12). 4. Raw data is consisted of green (LysM-GFP mice) and red (Texas Red dextran), but colors of each channel can be changed for analysis by selecting “Change colors” from the “Tools menu.” By comparing the intensity of red (S2) where blood leakage occurred and that (S1) of sites with no blood leakage (Fig. 3a), confirmation of vascular rupture at specific sites is possible by detecting an increase in the red intensity, indicating blood flow (Fig. 3b, c). 5. Select the area to be measured with the “ROI tool.” 6. Set a region of interest (ROI) using the “Rectangular ROI tool” (S1, S2) (see Note 13). 7. Click on the “Measurement item” at the top, which allows the measurement window to change. 8. Choose “Thresholding” from “Measurement protocol tasks.”

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Fig. 3 Analysis of vascular permeability in images obtained by two-photon intravital imaging. (a) Blood vessels (red) were labeled by dextran, and neutrophils (green) appeared on the image. Sites without leakage were designated as “S1,” the baseline intensity measurement. Sites of blood leakage were designated as “S2.” Scale bar: 30 μm. (b) The figure shows blood leakage over time in S2. Vessels and blood flow appear in magenta and rainbow colors, respectively. Color change indicates the amount of blood leakage. Scale bar: 10 μm. (c) When leakage occurs at S1 and S2, the increase in red intensity due to the blood outflow can be graphed. (Reproduced from Ref. [15] with permission from Hindawi)

9. Select “Find object Using % intensity” from “Thresholding”. 10. Choose “Channel” indicating blood flow, modify intensity, and set the following subcategories: (a) Drag “Exclude Not Touching ROIs” from “Filtering” to “Find object Using % intensity.” (b) Drag “Exclude touching Edge of Image” from “Filtering” to “Find object Using % intensity.” 11. When the adjustments are complete, select “Make Measurement Item” from the “Measurements menu,” and the data file will be added to the library. 12. The new data file obtained through analysis is summarized by the following items. Select “Analysis” next to “Display window (ROI)” and select “Data analysis.” 13. Finally, select “Edit Analysis” for a summary of data that suits research purposes. 14. Summarized data can be moved to a Prism software and displayed as a graph (see Note 14).

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Notes 1. Adequate size of imaging chamber is 90 mm (width) × 120 mm (length) × 25 mm (height). 2. Prepare a pair of thick plastic sheet with 65 mm (length) × 15 mm (height) and 45 mm (length) × 15 mm (height). 3. Mixture of SYLGARD 184 silicone elastomer curing agent and base should be fully mixed for even solidification. When mixing the curing agent and base, air bubbles might be formed. For

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rigidity of the silicone, air bubbles must be removed by applying vacuum using a desiccator. 4. Height of pedestal should slightly be lower than the dam. During cremaster muscle exposure, water flows into the chamber to prevent overflow. 5. If the molecular weight of dextran is too small, leakage from the blood vessel shortly after injection might occur. To avoid this possibility, high molecular weight dextran is recommended. Dextran of around 70 kDa has a molecular weight similar to that of albumin and is ideal for vascular permeability measurement. 6. Tunable Ti-Sapphire laser (690–1040 nm) is used as a light source for the LSM 7MP. Emission filter information; DAPI/ SHG, 420–480 nm; GFP/Alexa 488, 500–550 nm; and RFP/Alexa 555, 575–610 nm. 7. Residual fur from surgery might be detected while imaging due to strong autofluorescence. Hair removal cream may be used on mice fur after adjusting treatment times. 8. Precut sterile insect pins should be placed on both sides of muscle alternately in order to adequately stretch the muscles. 9. Since fluorochrome conjugated dextran solutions do not block the blood vessel, retro-orbital sinus vein injection may be easier in dextran visualization than tail vein injection. 10. During two-photon imaging, high laser power might induce cremaster muscle contraction, causing x, y-axis movement. 11. If the library already exists, it can be loaded by selecting “Open” from the “File menu.” The library is automatically saved when the library is closed. 12. A suitable filter only removes certain noises, but using an excessively strong filter may cause desired signals to disappear. 13. ROI tools can be selected depending on the shape of area to be analyzed. There are three ROI tools: “Rectangular ROI tool,” “Freehand ROI tool,” and “line tool line.” 14. Vascular permeability can be measured using an arbitrary unit, a.u. Blood vessel with no detectable leakage is set as the baseline for this unit.

Acknowledgments This study was supported by the National Research Foundation of Korea (NRF) grant funded by the Korea government (MSIT) (NRF-2019R1A2C2008481, Y-M.H.) Republic of Korea.

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References 1. Parnell E, Smith BO, Palmer TM, Terrin A, Zaccolo M, Yarwood SJ (2012) Regulation of the inflammatory response of vascular endothelial cells by EPAC1. Br J Pharmacol 166(2): 434–446. https://doi.org/10.1111/j. 1476-5381.2011.01808.x 2. Baskurt OK, Yalcin O, Meiselman HJ (2004) Hemorheology and vascular control mechanisms. Clin Hemorheol Microcirc 30(3–4): 169–178 3. Cueni LN, Detmar M (2008) The lymphatic system in health and disease. Lymphat Res Biol 6(3–4):109–122. https://doi.org/10.1089/ lrb.2008.1008 4. Vestweber D, Wessel F, Nottebaum AF (2014) Similarities and differences in the regulation of leukocyte extravasation and vascular permeability. Semin Immunopathol 36(2):177–192. https://doi.org/10.1007/s00281-0140419-7 5. Vestweber D (2012) Relevance of endothelial junctions in leukocyte extravasation and vascular permeability. Ann NY Acad Sci 1257:184– 192. https://doi.org/10.1111/j.1749-6632. 2012.06558.x 6. Phillipson M, Kubes P (2011) The neutrophil in vascular inflammation. Nat Med 17(11): 1381–1390. https://doi.org/10.1038/nm. 2514 7. Nourshargh S, Hordijk PL, Sixt M (2010) Breaching multiple barriers: leukocyte motility through venular walls and the interstitium. Nat Rev Mol Cell Biol 11(5):366–378. https:// doi.org/10.1038/nrm2889 8. Lanitis E, Irving M, Coukos G (2015) Targeting the tumor vasculature to enhance T cell

activity. Curr Opin Immunol 33:55–63. https://doi.org/10.1016/j.coi.2015.01.011 9. McDonald DM, Baluk P (2005) Imaging of angiogenesis in inflamed airways and tumors: newly formed blood vessels are not alike and may be wildly abnormal: Parker B. Francis Lecture Chest 128(6 Suppl):602S–608S. https:// doi.org/10.1378/chest.128.6_suppl.602S-a 10. Azzi S, Hebda JK, Gavard J (2013) Vascular permeability and drug delivery in cancers. Front Oncol 3:211. https://doi.org/10. 3389/fonc.2013.00211 11. Weis SM (2008) Vascular permeability in cardiovascular disease and cancer. Curr Opin Hematol 15(3):243–249. https://doi.org/ 10.1097/MOH.0b013e3282f97d86 12. Bates DO (2010) Vascular endothelial growth factors and vascular permeability. Cardiovasc Res 87(2):262–271. https://doi.org/10. 1093/cvr/cvq105 13. Sedgwick JB, Menon I, Gern JE, Busse WW (2002) Effects of inflammatory cytokines on the permeability of human lung microvascular endothelial cell monolayers and differential eosinophil transmigration. J Allergy Clin Immun 110(5):752–756. https://doi.org/ 10.1067/mai.2002.128581 14. Jaffer H, Adjei IM, Labhasetwar V (2013) Optical imaging to map blood-brain barrier leakage. Sci Rep UK 3. ARTN 3117. https:// doi.org/10.1038/srep03117 15. Park SA, Jeong S, Choe YH, Hyun YM (2018) Sensing of vascular permeability in inflamed vessel of live animal. J Anal Methods Chem 2018:5797152. https://doi.org/10.1155/ 2018/5797152

Chapter 3 Lymphatic Vascular Permeability Determined from Direct Measurements of Solute Flux Melanie Jannaway and Joshua P. Scallan Abstract The permeability of the lymphatic vasculature is tightly regulated to prevent the excessive leakage of lymph into the tissues, which has profound consequences for edema, immune responses, and lipid absorption. Dysregulated lymphatic permeability is associated with several diseases, including life-threatening chylothorax and pleural effusion that occur in patients with congenital lymphedema and lymphatic malformations. Due to a growing interest in uncovering new mechanisms regulating lymphatic vascular permeability, we recently pioneered methods to quantify this aspect of lymphatic function. Here, we detail our ex vivo method to determine the permeability of mouse collecting lymphatic vessels from direct measurements of solute flux. This method is modified from a similar ex vivo assay that we described for studying the contractile function of murine collecting lymphatic vessels. Since this method also uses the mouse as a model, it enables powerful genetic tools to be combined with this physiological assay to investigate signaling pathways regulating lymphatic vascular permeability. Key words Integrity, Leakage, Barrier function, Cannulation, Isolated vessel, Pressure, Perfusion, Endothelium

1

Introduction Microperfusion techniques have been used to study vascular permeability ever since Eugene Landis first developed his microinjection technique [1] to measure the water permeability of blood capillaries [2]. The Landis technique was later modified to include the cannulation of capillaries in vivo by a sharpened glass micropipette to control the osmotic and hydrostatic pressure of individual capillaries for the measurement of hydraulic conductivity [3]. While sensitive to small changes in endothelial barrier function, this approach required data from several hydrostatic pressures over a relatively long time to obtain a single data point. To determine the capillary permeability to various solutes over mere seconds, a quantitative solute permeability method was pioneered by Huxley and colleagues [4], which later utilized fluorescent dyes bound to

Binu Tharakan (ed.), Vascular Hyperpermeability: Methods and Protocols, Methods in Molecular Biology, vol. 2711, https://doi.org/10.1007/978-1-0716-3429-5_3, © Springer Science+Business Media, LLC, part of Springer Nature 2024

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albumin [5]. This method was later modified to allow the study of blood vessels in tissue beds that did not permit in vivo cannulation by surgically excising and cannulating the vessels on glass micropipettes [6]. Similarly, lymphatic vessels in the mouse are covered by adipocytes and extracellular matrix that hinders facile cannulation in vivo. To solve this problem, we have devised the ex vivo approach described here for quantifying solute flux across the murine lymphatic vessel wall. We recently described an assay that enabled the study of lymphatic contractile function using single popliteal collecting lymphatics surgically excised from the mouse [7]. However, we have updated various aspects of the method and combined it with quantitative fluorescence microscopy for investigating lymphatic vascular permeability. After surgical excision and cannulation of a collecting lymphatic vessel from the mouse mesentery, we were able to directly measure the solute flux to various fluorescent molecules and proteins. Using a dual photometer that splits fluorescence emission between green and red emitted wavelengths, we were able to determine the permeability to two solutes simultaneously [8].

2

Materials

2.1 Krebs Buffer and Fluorescent Tracers

1. Krebs buffer is made fresh weekly and contains (mM): 141.4 NaCl, 4.7 KCl, 2 CaCl2·2H2O, 1.2 MgSO4, 1.2 NaH2PO4·H2O, 3 NaHCO3, 1.5 Na-Hepes, and 5 D-glucose, diluted in ddH2O. Add 1 g/L BSA and adjust the pH to 7.4. 2. Aliquots of BSA dissolved in Krebs buffer and concentrated to >200 mg/mL using centrifugal filters with a MW cutoff of 30 kDa. 3. BSA conjugated to Alexa Fluor dyes (e.g., Alexa Fluor 488) according to the manufacturer’s protocol, aliquoted, and stored at -80 °C. Thaw and dilute this tracer on the day of use. 4. Other fluorescent tracers may be used as well (see Note 1).

2.2 Forceps and Spring Scissors

1. Fine scissors and extra fine Graefe forceps (Fine Science Tools, #11150-10 or #11152-10) are used to make a midline incision through which the mesentery is gently exteriorized using cotton-tipped applicators. 2. Curved, fenestrated ultra-fine Moria forceps are handsharpened under 40× magnification (see Note 2) and used for manipulating tissues in vivo and ex vivo. An extra pair is used for coarse dissection and is not sharpened. 3. Vannas-Tu¨bingen spring scissors are used for coarse dissection in vivo.

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4. Ultra-fine Moria spring scissors or McPherson-Vannas spring scissors are hand-sharpened under 40× magnification (see Note 2) to thin and reprofile the blades to make very small cuts. These are used for fine, controlled dissection of tissues ex vivo. 5. Dumont #5/45 forceps are used for the cannulation of single lymphatic vessels in a small chamber and for tying short silk sutures. 2.3 Dissection Chamber, Recessed into Table

1. Acrylic dissection chamber (see Note 3) that can be recessed into a table (see Fig. 1) should be used to permit the operator to work at the level of the vessel and rest their arms on the bench

Fig. 1 Acrylic dissection chamber. (a) The vessel is pinned into a custommachined acrylic dissection chamber when removing the connective tissue from the lymphatic vessel. (b) Importantly, the chamber is designed so that it can be recessed into a hole in the workbench, allowing the operator to work at the level of the vessel while keeping the forearms steady

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for stability. The bottom of the dissection chamber should be filled with 3–4 mm of black Sylgard 170. 2. 40 μm steel wire (Danish Myo Technology, #400447), cut to ~2 mm pieces, is used to pin the vessel onto the Sylgard. 3. Glass Pasteur pipette with a fire-polished tip is used to transfer the vessel from the dissection chamber into the cannulation chamber. The inside of the Pasteur pipette is siliconized (SurfaSil Siliconizing Fluid, #TS42800) to prevent the vessel from adhering to the glass surface. 4. An acrylic dissection board and a Sylgard dissection pedestal are also required for initial excision of the vessel from the anaesthetized mouse. 5. 4–0 black silk suture to be teased apart into individual strands 2–3 mm long (Roboz, #SUT-15-2) (see Note 4). 2.4 Glass Micropipette Fabrication

1. Borosilicate capillary glass for single lumen (World Precision Instruments, #TW150-6) and theta (World Precision Instruments, #TST150-6) micropipettes. 2. Rotary tool (Dremel) for grinding a hole in one side of the theta micropipette (see Note 5). 3. UV-curing optical cement (Norland Optical Adhesive, #NOA73) is used to make a plug behind the hole in the theta micropipette. 4. Pipette puller. 5. Microforge (Narishige, #MF-900) (see Note 6). 6. Brass rod handle (World Precision Instruments, #5444) to attach to each micropipette holder. 7. Micropipette holders (see Note 7) are custom-machined from acrylic rod with side ports that can mate to PE tubing (see Fig. 2).

2.5 Syringes for Filling Micropipettes and Controlling Pressure

1. 3 mL and 10 mL syringes (see Note 8). 2. Low protein binding PVDF filters, 0.45 μm. 3. Three-way stopcocks (Cole Parmer, #30600-09). 4. Blunt-ended hub adapters to mate stopcocks to PE-tubing (Component Supply Company, #05406-01). 5. Polyethylene tubing, cut to 14-inch pieces (Intramedic, PE-205).

2.6 Micropipette Mounting Stage and Pump

1. A breadboard (Thorlabs) was ordered in a custom size (16″ × 8″ × 0.5″; L × W × H) and used as a micropipette mounting stage. A 2″ diameter center hole was made that was later widened to a 3″ hole to make a lip at the bottom (see Fig. 3).

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Fig. 2 Micropipette holder on brass rod handle. (a) Micropipette holders are custom-machined from acrylic rod, which are then attached to a brass rod handle. (b) The acrylic micropipette holder has three components which screw together with O-rings at each connection. These O-rings are important for preventing leakage of pressure between each lumen (see Subheading 3.3). Side ports allow PE-205 tubing to be connected to the micropipette holder for back filling of the micropipettes via silastic tubing

Fig. 3 Mounting stage with water-jacketed cannulation chamber. (a) A custom-sized breadboard is used as the base of the mounting stage onto which two micromanipulators screw. (b) Overhead view of mounting stage with micromanipulators and inset cannulation chamber. (c) A hole is made in the center of the mounting stage, into which the cannulation chamber inserts. A lip on the bottom prevents the cannulation chamber from falling through the mounting stage, and a detent screw secures the chamber into the stage

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Fig. 4 Acrylic water-jacketed cannulation chamber. (a) Overhead view of the cannulation chamber, which is custom-machined from acrylic. The chamber is hollowed so that it is water-jacketed, with two ports connected to tubing that connects to the heat exchange pump (b). A glass coverslip is glued to the bottom of the cannulation chamber which holds the Krebs buffer within which the pipette tips are suspended, and the vessel is cannulated

2. Two 3-axis manual micromanipulators (Scientifica, #LBM-7) are attached to each end of the stage with ¼-20 threaded screws inserted into the threaded holes. 3. Round acrylic water-jacketed cannulation chamber (custommachined) fits in the 3″ center hole and is tightened with a detent screw (see Fig. 4). The lip on the bottom prevents it from falling through the hole. A coverslip is glued to the bottom of the cannulation chamber with flowable silicone sealant (Dow Corning, 734 RTV Sealant). Two nickel-plated brass barb adapters screw into the top of the cannulation chamber and connect to tubing from a heat exchanger pump to allow warm water to circulate within the chamber (see Note 9). 4. Heat exchanger pump (Lauda Alpha A6, #LCB4733-160002). 2.7 Dissecting Microscope and Light Source

1. Stereo dissection microscope with Greenough optics. The dissecting microscope should have 0.8–5× zoom at the very least. 2. Heavy duty dual arm boom stand with baseplate to stabilize the dissection microscope. 3. LED light source with gooseneck light guides for illuminating the dissection chamber (Techniquip ProLux LED Illuminator, #PLDM-X6X-TQB).

2.8 Inverted Fluorescence Microscope

1. Inverted microscope capable of fluorescence imaging (Zeiss Observer 7) should be outfitted with a sensitive CMOS camera (e.g., Hamamatsu Flash 4.0) and either a single or dual photometer (Horiba/PTI, #D-104).

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2. An LED light source permits excitation of fluorescent tracers as well as fluorescent proteins. To limit the emission light from overexciting the photomultiplier tubes, a 0.1% transmission neutral density filter is inserted into the light path (Chroma, #UVND 3.0) with a slider (see Note 10). 3. Windows computer running software for obtaining images and video from the microscope (Zeiss, Zen Blue). 2.9 Pressure Control System and Peristaltic Pump

1. Elveflow OB-1 pressure controller with three pressure ports (Elvesys, OB-1) (see Note 11). 2. Three low-pressure transducers. 3. Three analog amplifiers (Omega Engineering Inc., DP41-E-A) wired with BNC outputs. 4. An analog-to-digital interface with BNC terminals (National Instruments DAQ Device, USB-6341) for acquiring data from the amplifiers and photomultiplier tubes. 5. LabVIEW full development software for Windows (National Instruments, #776670-35). 6. Custom-written LabVIEW program for capturing data from the low-pressure transducers and photometer, as well as for controlling pressure through the OB-1. 7. Peristaltic pump (Gilson Minipuls 3, #61010-1-12 third Ed) (see Note 12).

3

Methods All procedures are performed at room temperature unless stated otherwise. Surgical tools and supplies need not be sterile for this terminal procedure. For the following protocol, collecting lymphatic vessels are harvested from the mesentery because lymphatic vessels from this tissue lack robust contractile function [9]. However, collecting lymphatic vessels from other tissues can be used to assess permeability if L-type Ca2+ channels are inhibited, which completely prevents spontaneous contractions [10].

3.1 Locate and Remove Mesenteric Lymphatic Vessel

1. Anesthetize a 6–8-week-old mouse with an i.p. injection of ketamine and an i.p. injection of thiobutabarbital (Inactin). Once anesthetized, shave the ventral abdomen of the mouse and place in supine position on an acrylic dissection board. Make a 2 cm midline incision through the skin and the abdominal wall. 2. Turn the mouse onto its left side and exteriorize the mesentery onto a clean Sylgard pedestal (0.5 cm thick) using cottontipped applicators that are moistened with Krebs buffer.

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Using a transfer pipette, moisten the mesentery with a few drops of Krebs buffer, and ensure that the tissue remains moistened throughout the procedure. 3. Under a dissection microscope, use moistened cotton swabs to gently maneuver the gut so that the mesentery lies flat on the Sylgard. Locate a collecting lymphatic vessel (see Note 13). 4. Grasp the adipose tissue or mesentery adjacent to the collecting lymphatic vessel with the coarse curved Moria forceps (see Note 14). Gently pull the tissue to the left, while the Vannas-Tu¨bingen spring scissors are used to cut a straight line parallel to the vessel on the left side. Then make an incision across the distal part of the vessel, taking care not to cut the artery or vein. Grasp the adipose tissue on the distal end of the vessel, and gently pull to the left again while cutting on the right side of the vessel, between the lymphatic vessel and the blood vessels. Once the desired length of vessel has been isolated, cut the proximal end of the lymphatic vessel to free it from the mesentery and immediately place it in a petri dish filled with Krebs buffer. 5. Once vessel collection is complete (see Note 15), euthanize the mouse according to your institutional protocol. 3.2 Dissect the Vessel from Connective Tissues

1. Fill the dissection chamber with Krebs buffer. Use the coarse curved Moria forceps to transfer two 40 μm steel pins to the dissection chamber and insert them into the Sylgard. Use the same forceps to transfer a lymphatic vessel segment to the dissection chamber. 2. Under the dissection microscope, push the pins into the adipose or connective tissue on each end of the lymphatic vessel (see Note 16) so that it lays flat on the Sylgard at approximately the same length as it was in vivo. 3. Using the sharpened fine-curved Moria forceps and the ultrafine spring scissors, grasp one end of the adipose tissue and gently pull to the left, while using the scissors to make very small cuts to remove strips of adipose tissue (see Note 17). Repeat cutting strips of tissue from the lymphatic vessel, moving closer to the lymphatic vessel wall. Alternate cutting with gentle teasing. As more tissue is cut from the vessel, it becomes easier to unravel the remaining adipose and connective tissue and resume cutting the tissue off of the vessel. Never pull the adipose or connective tissues off of the vessel (see Note 18). Once the adipose and connective tissues have been removed sufficiently for the vessel to sink in the Krebs buffer, move to the next step.

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Fig. 5 PE-205 tubing connects the syringe to the micropipette holder. A syringe connected to PE-205 tubing via a filter, a three-way stopcock, and hub adaptor attaches to the micropipette holder via a Luer port. This permits the back filling of the micropipettes with Krebs or tracer

3.3 Set Up the Micropipettes on the Mounting Stage

1. Fill the 10 mL syringe with Krebs buffer. Attach a 0.45 μm filter to the syringe. Attach a three-way stopcock to the filter. Attach a blunt-ended hub adapter (for PE-205 tubing) onto the threeway stopcock. Cut and attach a piece of PE-205 to the hub adapter. Use this syringe to fill the cannulation chamber on the stage with ~3 mL of filtered Krebs buffer. Then attach the end of the PE-205 to the output micropipette holder (see Fig. 5). 2. Insert the output (single lumen) micropipette into the holder, tighten the O-ring, and then use the syringe to backfill the micropipette. Lower the micropipette into the Krebs buffer, making sure there are no bubbles in the tip of the micropipette. 3. Fill a beaker or vial with 2 mL of Krebs buffer and add the appropriate amount of highly concentrated BSA to bring the concentration to 1% BSA. This is the washout buffer. In a second beaker or vial of 2 mL of Krebs buffer, add 1 mg of fluorescent BSA and the appropriate amount of unlabeled, concentrated BSA to bring the total concentration of BSA to 1%. 4. Fill two 3 mL syringes with the 1% BSA solutions from the beakers or vials. Then attach a 0.45 μm filter to each syringe. Attach a three-way stopcock to the filter. Attach the bluntended hub adapter to the three-way stopcock and affix a piece of PE-205 tubing.

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5. To mount and fill the input (theta) pipette, attach the PE-205 tubing from the 3 mL syringe containing the unlabeled BSA (washout buffer) to the micropipette holder. Fill one side of the theta micropipette up to the first O-ring and then carefully insert the micropipette. Tighten the O-ring, and then use the syringe to backfill the micropipette. 6. Attach the other 3 mL syringe to the other port on the micropipette holder and fill the holder with the fluorescent tracer. Tighten the second O-ring, and then use the syringe to fill the other half of the theta micropipette with fluorescent tracer. Lower the micropipette into the Krebs buffer (see Note 19). 7. Transfer the microscope. 3.4 Cannulate the Vessel

cannulation

chamber

to

the

dissecting

1. Before beginning cannulation, it is essential that the three-way stopcocks on all three syringes are open to the atmosphere; otherwise, the vessel will be irreversibly damaged and have to be discarded. 2. Place a 1 cm square piece of wet Kimwipe adjacent to or on the cannulation chamber. Under the dissecting microscope, tie a piece of suture into a loose single overhand knot. Do not tighten the knot. Repeat with another piece of suture. Transfer the two pretied sutures to the Krebs buffer in the cannulation chamber, and then remove the wet Kimwipe. Tie each suture very loosely onto the tip of each micropipette so that it stays but can be easily removed. Then lower the micropipettes toward the glass bottom of the cannulation chamber until the sutures make contact with the bottom. 3. Use a siliconized glass Pasteur pipette to transfer the vessel from the dissection chamber to the cannulation chamber (prefilled with Krebs). 4. Make sure that all syringes are sitting at the same height as the cannulation chamber (i.e., a pressure of ~1 cmH2O). 5. Use two Dumont 45° cannulation forceps to cannulate each end of the vessel onto the micropipettes. Try to identify the upstream side of the vessel if it contains a valve. Grab each side of this end with the forceps, gently pull each side apart to try to open the end of the vessel, and at the same time push the open end onto the micropipette. It is imperative that the forceps are grasping the very end of the vessel and not adipose or transparent elastic connective tissue (see Note 20). Once the vessel is on the micropipette, loosen the suture and move it over the vessel. Tighten the suture by applying even force on each side of the suture so that it secures the vessel to the micropipette. 6. Once the upstream side of the vessel is cannulated onto the input (theta) micropipette, carefully raise the two 3 mL

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syringes by ~5 cm to pressurize the lymphatic vessel. If the vessel distends easily, then it has been successfully cannulated permitting flow from both sides of the theta micropipette. This step will also remove any twists in the vessel and make it easier to cannulate the downstream (output) end of the vessel. 7. Repeat step 5 to cannulate the other end of the lymphatic vessel onto the output (single-lumen) micropipette. With practice, the cannulation chamber should not have to be rotated to cannulate this side. To verify that the output side has been cannulated and permits flow through the vessel, raise the 10 mL syringe by ~5 cm. The vessel should easily become distended. Alternatively, gentle tapping on the PE tubing should cause any valve leaflets present to flutter easily. 3.5 Move the Cannulation Chamber to the Microscope

1. Transfer the isolated vessel apparatus to the microscope stage. While previous versions of an isolated vessel stage had micromanipulators that hung below microscope stage [7], thus limiting the number of microscope stages it could be used with, the isolated vessel stage described here overcomes these previous limitations and can be used on any microscope stage. The isolated vessel apparatus we use does not secure to the microscope stage but has sufficient mass to be stable. 2. Slowly and carefully disconnect the PE tubing from the bluntended hub adapters on the syringes making sure not to pinch or rapidly move the tubing (see Note 21). Attach the PE tubing to the pressure control system with silastic tubing that makes a tight seal around PE-205 tubing. The pressure system should be set at 3 cmH2O for all pressures prior to connecting the PE tubing. Once the tubing is connected, change the input pressures to 10 cmH2O and output pressure to 8 cmH2O to permit flow that primes the vessel with fluorescent tracer and washout buffer and prevents their mixing (see Note 22). 3. Place a container of Krebs buffer near the peristaltic pump. Connect the tubing from the peristaltic pump to the cannulation chamber so that it slowly adds the Krebs buffer to the vessel at a rate of 0.5 mL/min. Attach a vacuum line on the opposite side of the chamber to remove excess Krebs buffer at a constant rate. The volume of the Krebs buffer will affect temperature, so the vacuum should not continuously remove fluid. 4. Connect the tubing from the heat exchanger pump to the cannulation chamber and turn on the pump. This will need to be calibrated prior to the experiment to ensure that the heated water jacket will warm the Krebs buffer to 37 °C over the course of ~1 h. 5. After equilibrating the Krebs buffer to 37 °C (verify with a thermometer) for 1 h, the experiment may begin.

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3.6 Conduct the Experiment

1. The simplest protocol to perform is to measure the solute flux of a wild-type lymphatic vessel at multiple pressures. First, set the two theta micropipette pressures to be equal and change the light path to use the camera mounted on the microscope to monitor the flow of each side. The flow of the fluorescent tracer and washout buffer should be ~50:50 at equal pressures. Reduce the pressure of the washout side until the fluorescent tracer fills the lumen of the vessel and just starts to enter the washout side. Reset the input pressures to be equal again and then lower the pressure of the fluorescent tracer side until only a small amount of fluorescent tracer is streaming through the lumen (this prevents mixing of the washout buffer and fluorescent tracer). The use of software that enables instantaneous switching between these two pairs of pressures is highly recommended because this allows for rapid switching between complete perfusion of the fluorescent tracer or the washout buffer. 2. Once the input pressures are determined for a given output pressure, perfuse the washout buffer to make sure there is no visible fluorescence in the field of view. 3. Set a region of interest (ROI) using the metal leaflets on the photometer to obtain photons from a rectangular area encompassing the vessel lumen and bath solution on either side of the vessel. 4. Excite the fluorescent tracer within the vessel with the LED. Attenuate the excitation light intensity by sliding the neutral density filter into the light path. Change the light path to send the emitted fluorescence light to the photometer and begin recording light intensity. 5. After collecting data for a stable baseline, use the software to switch the pressures to fill the vessel lumen with fluorescent tracer, which will cause a step increase in photometer voltage. Over time, the photometer voltage will gradually increase in a linear fashion, generating a line with a defined slope as solute moves out of the vessel and accumulates in the surrounding bath solution. 6. Once a steady, linear slope has been obtained, switch the pressures to wash out the fluorescence, returning the photometer voltage back to baseline. 7. Take a brightfield image of the vessel at this pressure in Zen and use the Distance tool to measure the internal diameter of the vessel. 8. Repeat steps 1–8 for each pressure of interest without changing the ROI.

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9. The permeability of the vessel can now be calculated for each pressure according to a modified form of Fick’s first law using the raw trace and the diameter of the vessel as previously published [4, 8, 11, 12]. 3.7 Cleaning of Equipment

1. Set pressures for all lumens to 3 cmH2O. Detach the isolated vessel apparatus from the pressure system. 2. The micropipettes can be reused if cleaned well. Untie the sutures to remove the vessel, and then retie the sutures onto the ends of each micropipette for future use. Remove the output micropipette from the holder, and attach a vacuum line to the rear and dip in ddH2O 10 times, filling the micropipette at least halfway each time. Do the same with acetone five times. Store the micropipette in a covered tray to protect from dust. 3. To clean the theta micropipette, we use a spare micropipette holder attached to the vacuum line to simultaneously clean both sides of the theta micropipette as described above. 4. The pipette holders should be rinsed thoroughly with ddH2O. 5. Remove the cannulation chamber from the mounting stage and clean with 10% Contrad detergent and a cotton-tipped applicator. Rinse with ddH2O, then 30% ethanol, and ddH2O again. Blow dry immediately with pressurized air, before returning it to the mounting stage. 6. Rinse the dissection chamber with warm ddH2O twice, followed by 70% ethanol. Use a vacuum line to remove each solution, and then allow to air dry overnight. 7. Clean the acrylic dissection board and Sylgard pedestal with 10% Contrad detergent and rinse with ddH2O, before allowing to air dry. 8. The three-way stopcocks are rinsed with 10% Contrad detergent, then ddH2O, then 35% ethanol, and again with ddH2O. All are then left to air dry. 9. All dissection forceps and scissors should be very carefully cleaned with ddH2O and then 70% ethanol on a gauze pad and then returned to their storage cases.

4

Notes 1. The measurement of solute flux is achieved by perfusing a fluorescent tracer or combination of two tracers with nonoverlapping fluorescence emissions. A variety of fluorescent tracers can be used including, but not limited to, sodium fluorescein, Alexa Fluor-conjugated BSA; Alexa Fluor-conjugated

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α-lactalbumin [4], or Alexa Fluor-conjugated dextrans. The use of FITC-conjugated tracers is not recommended because FITC generates reactive oxygen species which will alter solute permeability [13]. The simultaneous measurement of two tracers is also possible (see Note 10); we have found that solutes conjugated to Alexa Fluor 488 and Alexa Fluor 594 can be used together. 2. Aluminum oxide lapping film adhesive sheets (3M, Amazon) are affixed to one side of a glass slide and used to sharpen the fine forceps and spring scissors while under 40× magnification under a dissecting microscope. Specifically, tools are sharpened with 12 μm (yellow, 1200 grit) paper, followed by 3 μm (pink, 8000 grit) paper, and polished with 0.3 μm (white, 60,000 grit) paper. Sometimes, damaged tools first need to be reprofiled with 30 μm (green, 600 grit) paper. 3. Our acrylic dissection chamber is custom-machined, and its dimensions are 4.5 cm in height, 10 cm diameter excluding lip, 12.5 cm diameter including lip, and a 5 cm bath diameter. 4. Tie a single overhand knot using one piece of silk suture (2 mm in length) on a wet Kimwipe using the cannulation forceps. Each knot is then looped over each micropipette tip, where it is secured temporarily prior to vessel cannulation. 5. For the theta pipette only, use a rotary tool such as a Dremel to create a small 1 mm hole on one side of the pipette approximately 2 cm from the back of the pipette. Clean the pipette with acetone to remove any glass debris. Fill the pipette behind the hole with UV optical cement and expose it to a UV light for ~5 min to cure the cement (Ulako UV365nm, Amazon). Be careful to avoid letting cement drip into the side without the side hole. 6. Once the pipette has been pulled (length ~8 cm), use the microforge to cleanly break the end of both pipettes to the desired diameter. For mouse mesenteric lymphatic vessels, this is approximately 60 μm (outer diameter). Then, fire polish the tip of the pipette using the microforge, which ensures the vessel will be cannulated smoothly without damaging the endothelium. The tip of the micropipette is bent to a 30° angle using the microforge so that the micropipette will be parallel with the bottom of the cannulation chamber. 7. The micropipette holder on the input side has two separate chambers so that one side of the theta pipette can be filled with two different solutions. O-rings are placed inside the micropipette holders to create a tight seal around the pipettes and prevent leakage of pressure between the two lumens (World Precision Instruments Replacement Gasket, 1.5 mm, #GO3100).

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8. Each syringe is attached to a filter followed by a three-way stopcock, then a hub adapter which is connected to PE 205 tubing. For the input pipette, two 3 mL plastic syringes are used, one filled with Krebs buffer (washout) and one with Krebs buffer containing the tracer(s) of choice. For the output pipette, one 10 mL plastic syringe containing Krebs buffer is required. 9. The cannulation chamber on the mounting stage should have a water jacket to warm the lymphatic vessel up to physiological temperature (37 °C) before the experiment. Our chamber is custom machined, with a diameter of 3″ and height of 5/16″, and fits flush with the top of the stage. The chamber where the vessel will be submerged in Krebs buffer is 6.5 cm long, 9 mm at its narrowest and 19 mm at its widest. It needs to be wide enough to be able to comfortably fit and maneuver cannulation forceps. The bottom is made with a glass cover slip (Warner Cover Glass, #1 thickness, #CS-24/50). 10. To quantify the solute flux for two different fluorescent tracers simultaneously, the photometer must separate incoming light into two nonoverlapping wavelengths (e.g., green and red) with an appropriate dichroic filter (e.g., Chroma T565lpxr) mounted in a Nikon 18 mm cube (Chroma 91000). Then emission filters for each wavelength (e.g., Chroma ET525/ 50 m and Chroma ET645/75 m) must be installed immediately before each photomultiplier tube to further select for a single bandpass. 11. Alternatively, the simplest way to control pressure is to attach each micropipette lumen to a syringe filled with water and mounted to the wall. Then the syringe is connected to the micropipettes via PE tubing. Pressure is controlled by raising the syringes above the microscope stage, where pressure is zero. Since the syringes are filled with water, raising them by 1 cm creates 1 cmH2O pressure. 12. The Krebs buffer inside the cannulation chamber is continuously exchanged using a peristaltic pump. Exchange of Krebs is critical to prevent evaporation that will cause concentration of the salts and alter the pH of the solution, and barrier function will be negatively affected. 13. The mesenteric collecting lymphatic vessels are located parallel to each artery and vein pair, running from the lymph node to the intestinal border. In the duodenum and jejunum, mesenteric collecting lymphatics are usually white in appearance due to chyle transport after overnight feeding, have several valves that are apparent, and are distended. Otherwise, they are transparent in color and become difficult to locate for the novice.

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With experience, we are able to locate both types of collecting lymphatic vessels. 14. Avoid overstretching the vessels. Excessive axial or radial stretching of the lymphatic vessel will cause irreversible damage to the endothelium and smooth muscle layers and the vessel cannot be used for experiments. 15. It is recommended to remove several collecting lymphatic vessels from each mouse to increase n-number. This also limits the chances of obtaining collecting lymphatic vessels that have branches, which will leak fluorescent tracer at a very high rate, and also increases the chances of obtaining vessels with the desired number of valves. 16. Do not push the pins through the lymphatic vessel itself. Rather, gently push the pins through the associated connective tissue. 17. When removing the adipocytes from the vessel, work methodically from the outside in, slowly making cuts closer to the vessel wall. Always cut on the left-hand side of the vessel. Rotate the dissection dish to cut on the other side. Never cut or nick the vessel as this will lead to obvious fluorescent tracer leakage on the microscope and the vessel will need to be discarded. It is not necessary to remove all adipocytes, but the ends of the vessel need to be free of adipocytes to permit cannulation in the next step. 18. Each vessel should have the following baseline criteria to be used for an experiment: no branches, no holes, no leakage at cannulation sites, and no constricted spots indicating damaged smooth muscle cells. Depending on the experimental protocol in use, the vessel also requires the desired number of valves. 19. At this point, if the micropipettes are filled correctly, then fluorescent tracer should be seen slowly leaking out of the micropipette tip into the Krebs buffer inside the cannulation chamber. Further, if the input syringes are raised, fluorescent tracer can be observed leaking out at a higher rate. If tracer is not visualized leaking from the micropipettes, check the micropipette for any bubbles or debris. 20. The ends of the vessel will necessarily be damaged by the forceps; however, the sutures will be tied after the damaged ends of the vessel, so this will not contribute to any measurement of solute flux. There should be no visible leakage of fluorescent tracer from the ends of the vessel if the sutures are tied tight enough. The sutures should not be tied so tight that they cannot be removed after the experiment. Take care not to bend or kink the sutures as this creates a weak spot in the suture that can break. We routinely reuse the same pair of sutures for several experiments before having to retie new sutures.

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21. The PE tubing can only be connected or disconnected when the three-way stopcock valve is open to the atmosphere. If the tubing is connected or disconnected with the valve closed, then the vessel will be irreversibly damaged and must be discarded. 22. Mixing of washout buffer in the fluorescent tracer must be avoided; otherwise, the fluorescent tracer will be diluted to an unknown concentration, thus voiding the solute flux measurement. References 1. Landis EM (1926) The capillary pressure in frog mesentery as determined by microinjection methods. Am J Phys 75:548–570 2. Landis EM (1927) Micro-injection studies of capillary permeability. II. The relation between capillary pressure and the rate at which fluid passes through the walls of single capillaries. Am J Phys 82:217–238 3. Michel CC, Mason JC, Curry FE, Tooke JE, Hunter PJ (1974) A development of the Landis technique for measuring the filtration coefficient of individual capillaries in the frog mesentery. Q J Exp Physiol Cogn Med Sci 59(4):283–309. https://doi.org/10.1113/ expphysiol.1974.sp002275 4. Huxley VH, Curry FE, Adamson RH (1987) Quantitative fluorescence microscopy on single capillaries: alpha-lactalbumin transport. Am J Phys 252(1 Pt 2):H188–H197. https://doi. org/10.1152/ajpheart.1987.252.1.H188 5. Rumbaut RE, Huxley VH (2002) Similar permeability responses to nitric oxide synthase inhibitors of venules from three animal species. Microvasc Res 64(1):21–31. https://doi.org/ 10.1006/mvre.2002.2394 6. Yuan Y, Chilian WM, Granger HJ, Zawieja DC (1993) Permeability to albumin in isolated coronary venules. Am J Phys 265(2 Pt 2):H543– H552. https://doi.org/10.1152/ajpheart. 1993.265.2.H543 7. Castorena-Gonzalez JA, Scallan JP, Davis MJ (2018) Methods for assessing the contractile function of mouse lymphatic vessels ex vivo. Methods Mol Biol 1846:229–248. https:// doi.org/10.1007/978-1-4939-8712-2_15

8. Jannaway M, Scallan JP (2021) VE-cadherin and vesicles differentially regulate lymphatic vascular permeability to solutes of various sizes. Front Physiol 12:687563 9. Zawieja SD, Castorena-Gonzalez JA, Scallan JP, Davis MJ (2018) Differences in L-type Ca (2+) channel activity partially underlie the regional dichotomy in pumping behavior by murine peripheral and visceral lymphatic vessels. Am J Physiol Heart Circ Physiol 314(5): H991–H1010. https://doi.org/10.1152/ ajpheart.00499.2017 10. To KHT, Gui P, Li M, Zawieja SD, CastorenaGonzalez JA, Davis MJ (2020) T-type, but not L-type, voltage-gated calcium channels are dispensable for lymphatic pacemaking and spontaneous contractions. Sci Rep 10(1):70. https://doi.org/10.1038/s41598-01956953-3 11. Scallan JP, Hill MA, Davis MJ (2015) Lymphatic vascular integrity is disrupted in type 2 diabetes due to impaired nitric oxide signalling. Cardiovasc Res 107(1):89–97. https:// doi.org/10.1093/cvr/cvv117 12. Scallan JP, Huxley VH (2010) In vivo determination of collecting lymphatic vessel permeability to albumin: a role for lymphatics in exchange. J Physiol 588(Pt 1):243–254. https://doi.org/10.1113/jphysiol.2009. 179622 13. Bingaman S, Huxley VH, Rumbaut RE (2003) Fluorescent dyes modify properties of proteins used in microvascular research. Microcirculation 10(2):221–231. https://doi.org/10. 1038/sj.mn.7800186

Chapter 4 Evaluation of Mesenteric Microvascular Hyperpermeability Following Hemorrhagic Shock Using Intravital Microscopy Taylor R. Williams and Ed W. Childs Abstract Intravital microscopy is a powerful tool for evaluating vascular hyperpermeability in various vascular beds. Hemorrhagic shock after traumatic injury is known to induce microvascular hyperpermeability, lifethreatening edema, and microcirculatory perfusion disturbances. Here we describe the microsurgical and imaging methods to study mesenteric vascular hyperpermeability using intravital microscopy, in a rat model of hemorrhagic shock. In this protocol, hemorrhagic shock is induced by controlled withdrawal of blood to reduce the mean arterial pressure (MAP) to 40 mmHg for 60 min, followed by resuscitation for 60 min. To study the changes in vascular permeability, the rats are given FITC-albumin, a fluorescent tracer, intravenously. The FITC-albumin flux across the vessel wall is measured in mesenteric postcapillary venules by determining intravascular and extravascular fluorescence intensity under intravital microscopy. Intravital microscopic evaluation of high molecular weight FITC-albumin permeability is a reliable indicator of microvascular hyperpermeability. Key words Vascular hyperpermeability, Hemorrhagic shock, Intravital microscopy

1

Introduction Intravital microscopy is one of the most reliable methods for measuring vascular hyperpermeability in laboratory rodents in vivo. Understanding, minimizing, and/or reversing vascular hyperpermeability during shock would have a tremendous impact in preclinical research and the treatment of critically ill patients. The methods we describe here provide a means to study this phenomenon in a reproducible and efficacious manner in the laboratory. Using this technique, we can change variables (such as medications, hormones, genetics, etc.) and assess their effects on vascular permeability [1–5]. Vascular hyperpermeability and associated edema occur as a serious clinical complication of hemorrhagic shock (HS) and many other conditions. Hemorrhagic shock has been implicated in the pathogenesis of multiple organ failure and accounts for 30%

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of deaths associated with traumatic injury in affected patients [6]. Vascular hyperpermeability, the excessive leakage of fluids and proteins from the intravascular area to the interstitium, may help explain the mechanism behind the edema, microcirculatory perfusion disturbances, and multiorgan failures that occur following hemorrhagic shock [11]. The limitation to precisely determine this extravascular leakage of fluid and proteins is a limiting factor in many labs that conduct research in endothelial barrier functions and vascular permeability. Intravital microscopy using a fluorescent tracer is considered as the gold standard for measuring vascular hyperpermeability in research animal models in vivo. Vascular hyperpermeability following hemorrhagic shock has also been shown by several investigators to be associated with induction of oxidative stress, reactive oxygen species (ROS) formation, and the activation of apoptotic signaling pathways [6–9]. When oxygen is re-introduced into the ischemic tissue during reperfusion, a burst of ROS is subsequently formed and introduced as well. The ischemia-reperfusion process selectively damages endothelial cells and barriers through a mechanism that involves oxygenderived free radicals [9, 10]. Thus, oxidative stress due to ROS formation is a major trigger for vascular hyperpermeability and an important area of research [2]. Additionally, recent studies have demonstrated that the activation of the apoptotic signaling pathway is critical to inducing the vascular hyperpermeability associated with hemorrhagic shock. Thus, suggesting that the inhibition of this pathway would provide protection against vascular hyperpermeability in a rat model of hemorrhagic shock [4].

2

Materials

2.1 Anesthesia Induction

1. 50% urethane solution. 2. 28-Gauge insulin needles. 3. 1 mL syringe.

2.2 Operative Equipment

1. Rat (male 275–325 g). 2. Heat lamp. 3. Heated surgical table. 4. Absorbent under pad. 5. Microdissection curved forceps. 6. Microdissection scissors. 7. Small hemostat. 8. Lab tape. 9. 6-0 silk suture.

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10. Normal saline. 11. Heparinized saline. 12. Polyethylene tubing (“PE-50”; 0.58 mM internal diameter). 13. Three-way stop cocks × 3. 14. 23-gauge one-inch needles. 15. Micro-clip (“bulldog clamp”). 16. Large plexiglass plate. 17. Small plexiglass plate. 18. Cotton tip applicator. 19. 2 × 2 gauze. 20. Plastic wrap. 2.3

Shock Induction

1. Arterial blood pressure monitor. 2. 10 mL syringe × 2. 3. Syringe pump.

2.4 Vascular Permeability Monitoring

3 3.1

1. Intravital microscope with camera. 2. Computer with intravital microscopy software. 3. Fluorescein isothiocyanate conjugate – albumin, bovine.

Methods Anesthesia

1. Position the awake 275–325 g rat in the prone position on the surgical table. Make sure the thighs are exposed. 2. Inject of 50% urethane (0.35 mL/100 g of rat weight) intramuscularly into the thigh of the rat (see Note 1). 3. Place the rat back into the cage and allow anesthesia to take full effect (30 min to 1 h).

3.2

Preparation

1. Clip the fur on the rat’s neck, abdomen, and inner thighs (see Note 2). 2. Position the head of the rat toward the operator. 3. Place the rat in the supine position on the heated (43 °C) surgical table. The absorbent under pad should be placed between the rat and the surgical table. 4. Secure all four limbs using lab tape.

3.3

Jugular Vein

1. Grasp the skin of the right side of the neck just superior to the clavicle and using dissection scissors, cut a 1 cm opening through the skin.

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2. Grasp the underlying fascia and dissect through sharply with scissors. 3. Identify the underlying external jugular vein and bluntly dissect away the surrounding tissue. 4. Isolate the external jugular vein by passing a right-angled forceps with a 6-0 silk suture deep to the vein. 5. Tie the suture at the most cephalad portion of the vein. Clamp the suture to the rat’s jaw under slight tension. 6. Pass another 6-0 silk suture around the most caudal portion of the vein. Make a knot loop but do not tie it down. 7. Using the microdissection scissors, make a transverse cut to the anterior surface of the vein. 8. Connect a three-way stop cock to a cannula flushed with heparinized saline. Insert this cannula bevel side up into the vein. 9. Open the three-way stop cock and ensure back flow. 10. Once back flow is confirmed, tie the second (caudal) silk suture over the portion of the cannula within the vessel. 11. Unclamp the first (cephalad) silk suture from the rat’s jaw and tie it around the cannula, securing it to the vein. 3.4

Carotid Artery

1. Identify the midline of the neck through the incision. On either side will be the sternohyoid muscle. Lateral to the sternohyoids will be the sternocephalic muscles. Bluntly separate the sternohyoid and sternocephalic muscles to expose the carotid artery, lateral to the trachea. 2. Isolate the carotid artery with a right-angled forceps. Bluntly dissect away the surrounding tissue. 3. Pass the right-angled forceps deep to the vessel and isolate it by passing 6-0 silk suture around it. 4. Tie the first suture at the most cephalad portion of the artery. Clamp this suture to the rat’s jaw under slight tension. 5. Place a bulldog clamp on the inferior portion of the vessel. 6. Pass another 6-0 silk suture around the most caudal portion of the artery. Make a knot loop but do not tie it down. 7. Use the scissors to make a transverse arteriotomy on the anterior surface of the artery. 8. Connect a three-way stop cock to a cannula flushed with heparinized saline. Insert this cannula bevel side up into the vein. 9. Use the index finger of your nondominant hand to secure the cannula to the chin of the rat. Then remove the bulldog clamp. 10. While still securing the cannula, use your dominant hand to quickly open and close stop cock to ensure bleeding.

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11. Flush cannula with heparinized saline. 12. Tighten the inferior knot. Release superior knot from clamp on the chin and tie it around the cannula. 3.5

Femoral Artery

1. Turn the head of the rat away from the operator. 2. Identify the depression three quarters of the distance from the pubic tubercle to the inner border of the leg. 3. Grasp the overlying skin with forceps and use dissection scissors to cut a 1.5 cm circular opening. 4. Sharply cut through the underlying fascia using the scissors. 5. Identify the femoral vessels above the takeoff of the epigastric vessels. 6. Use the right-angled forceps to isolate the femoral artery and pass a 6-0 silk suture around artery. Tie this suture as caudally as possible, just above the level of the epigastric vessels. 7. Place a bulldog clamp at the superior portion of the artery. 8. Pass another 6-0 silk suture just below the clamp. Make a loop but do not tie it down. 9. Make a transverse arteriotomy and insert the stretched heparinized saline flushed cannula (see Note 3). 10. Loosely secure the superior suture. 11. Remove the clamp and flash the stop cock to ensure back bleeding. 12. Flush with heparinized saline and secure the suture.

3.6

Mesentery

1. Identify the midpoint between the xyphoid process and pubic tubercle. Use scissors to make a longitudinal midline incision extending 1 inch cephalically and 1 inch caudally. 2. Visualize the white line in the midline and incise the fascia sharply at this point. 3. Remove the tape from the limbs and gently reposition the rat onto its left side on a plexiglass plate. You can pick the rat up by its limbs to facilitate repositioning. 4. Place a smaller plexiglass plate in front of the rat’s abdomen. Apply saline to the surface of the plate. 5. Place your thumbs on the right lateral abdominal wall, and massage the abdominal contents out of the abdominal cavity. Apply pressure to the posterior surface of the spine to help facilitate this step. 6. Use a cotton-tip applicator to splay out the bowel and its mesentery onto the smaller plexiglass. 7. Identify a capillary within the clear mesentery (see Note 4).

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8. Use the cotton-tip applicator to carefully return all other portions of the bowel back into the abdominal cavity. 9. Place several normal saline soaked 2 × 2 gauze sponges over the bowel while leaving the mesentery and chosen capillary exposed (see Note 5). 10. Cut a piece of plastic wrap and lay it over the mesentery and small bowel. It should lie in contact with the mesentery (see Note 5). 3.7

Preshock

1. Allow a postoperative 30-min recovery period. After this time place the rat (still on plexiglass) on the microscope and visualize the exposed capillary through the microscope (Fig. 1). 2. Inject FITC-albumin (50 mg/kg) through the jugular catheter (see Note 8) (Fig. 1b). 3. Place an anal temperature probe. Adjust heat lamp as necessary to maintain temperature between 36 and 38 °C. 4. Connect the femoral artery stop cock to a blood pressure transducer. 5. Flush a 10 mL syringe with heparinized saline and then connect the empty syringe to the carotid artery catheter. 6. Connect the jugular catheter to a syringe pump filled with normal saline. Infuse at a rate of 3 mL/h. 7. Take a preshock microscopic image and save it for reference (see Note 6) (Fig. 1a, b).

3.8

Shock

1. Use the empty syringe connected to the carotid catheter to draw blood from the rat until the mean arterial pressure (MAP) is reduced to 40 mmHg. 2. Keep the MAP around 40 mmHg (38–42 mmHg) by drawing more or returning blood as necessary. 3. Continue this process for the desired period. We typically shock the rats for 60 min. 4. After the planned shock period, return the shed blood to the rat over a 5-min duration. 5. Supplement with normal saline as needed to maintain MAP above 90 mm Hg. 6. Take microscopy images at desired time intervals. We typically take images at 10-min intervals postshock (Fig. 1) (see Notes 6 and 7).

3.9 Intravital Microscopy

1. Open images using your intravital microscope software. 2. Analyze the fluorescence intensity at two locations (both inside and outside of the vessel) on the preshock control image (Tc). Average each and create an out-in ratio.

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A

Sham

(∆I) FITC Fluorescence Intensity

B

Shock T60 Shock Sham

4

3 Control 2

30 min.

*

Shock 60 min.

*

*

*

*

1

0

10

20

30

Start Resuscitation

Time (min.)

40

50

60

Tharakan et al, Cell Death Discovery, 1:15042, 2015

Fig. 1 Intravital microscopic imaging and evaluation of vascular hyperpermeability in a rat model of hemorrhagic shock. The images of mesenteric postcapillary venules from sham and hemorrhagic shock for 1 h followed by 60 min of resuscitation (label T60) (a). Permeability is expressed as change in fluorescence intensity inside the vessel compared to the intensity outside the vessel (b). FITC-albumin extravasation is significantly high following shock compared to sham group. * ( p < 0.05). The figure is taken and modified from our previously published study. (Tharakan et al., Cell Death Discovery,1:15042, 2015; Creative Commons License https://creativecommons.org/licenses/)

3. Repeat for the rest of the images (Tx) taken during the shock period. Use the same locations along the vessel as the control image. 4. Obtain a ratio (Tx:Tc) for each of the time points. 5. Plot and compare your ratios as desired.

4

Notes 1. Administer half of the anesthetic dose into each thigh for faster onset of anesthesia. 2. Ensure that the rats are fasted for 18 h prior to the procedure.

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3. Due to the smaller caliber of the femoral artery, stretch the PE-50 cannula at the tip in order to narrow it. 4. The capillaries in the mesentery are almost invisible to the untrained eye. You will need to look carefully for a very small vessel, ideally 25–30 microns in diameter. 5. The mesentery should always be kept moist with warm normal saline. The overlying plastic wrap will help reduce evaporation. 6. Images may be captured at desired intervals and selected short video clips may be saved as necessary. 7. The rat should be normothermic (~37°). Heat lamps may be necessary during surgery and during shock to maintain this temperature. 8. Turn lights off when handling the FITC-albumin. References 1. Childs E et al (1999) Leukocyte adherence and sequestration following hemorrhagic shock and total ischemia in rats. Shock 11:248–252 2. Childs EW, Udobi KF, Wood JG, Hunter FA, Smalley DM, Cheung LY (2002) In vivo visualization of reactive oxidants and leukocyteendothelial adherence following hemorrhagic shock. Shock 8:423–427 3. Tharakan B et al (2010) (-)-Deprenyl inhibits vascular hyperpermeability after hemorrhagic shock. Shock 33:56–63 4. Childs E et al (2007) Apoptotic signaling induces hyperpermeability following hemorrhagic shock. Am J Physiol Heart Circ Physiol 292:H3179–H3189 5. Childs E et al (2010) 17β-estradiol mediated protection against vascular leak after hemorrhagic shock: role of estrogen receptors and apoptotic signaling. Shock 34:229–235 6. Murao Y, Hata M, Ohnishi K, Okuchi K, Nakajima Y, Hiasa Y, Junger WG, Hoyt DB, Ohnishi T (2003) Hypertonic saline resuscitation reduces apoptosis and tissue damage of the small intestine in a mouse model of hemorrhagic shock. Shock 20:23–28 7. Childs EW, Udobi KF, Hunter FA, Dhevan V (2005) Evidence of transcellular albumin

transport after hemorrhagic shock. Shock 23: 565–570 8. Davidson MT, Deitch EA, Lu Q, Hasko G, Abungu B, Nemeth ZH, Zaets SB, Gaspers LD, Thomas AP, Xu DZ (2004) Traumahemorrhagic shock mesenteric lymph induces endothelial apoptosis that involves both caspase-dependent and caspase-independent mechanisms. Ann Surg 240:123–131 9. Therade-Matharan S, Laemmel E, Duranteau J, Vicaut E (2004) Reoxygenation after hypoxia and glucose depletion causes reactive oxygen species production by mitochondria in HUVEC. Am J Physiol Regul Integr Comp Physiol 287:R1037–R1043 10. Savoye G, Tamion F, Richard V, Varin R, Thuillez C (2005) Hemorrhagic shock resuscitation affects early and selective mesenteric artery endothelial function through a free radical-dependent mechanism. Shock 23:411– 416 11. van Leeuwen ALI et al (2020) In vitro endothelial hyperpermeability occurs early following traumatic hemorrhagic shock. Clin Hemorheol Microcirc 75:121–133

Chapter 5 Determination of Endothelial Barrier Resistance by Electric Cell-Substrate Impedance Sensing (ECIS) System Hemant Giri and Madhulika Dixit Abstract Endothelial cells lining the inner surface of blood vessels and lymphatic vessels play an indispensable role in vascular homeostasis. Apart from regulating vessel tone and forming an anti-thrombotic and antiatherosclerotic surface, the dynamic endothelial barrier controls transport of solutes and fluid in and out of tissues at the capillary bed. Transit of circulating leukocytes into and out of circulation during inflammation and tissue repair is also regulated by the endothelium. Dysregulation of this barrier function of endothelial cells is a hallmark feature of multiple diseases and conditions such as sepsis, cancer metastasis, and edema. In this chapter we describe a detailed methodology to perform an in vitro experiment to monitor changes in barrier properties of human umbilical vein endothelial cells (HUVECs) in real time, in response to thrombin with electrical cell-substrate impedance sensing (ECIS) biosensor system. Key words Vascular endothelial barrier, Endothelial barrier resistance, Electrical cell-substrate impedance sensing

1

Introduction Maintenance of endothelial barrier function is vital for physiology of all organ systems. These barriers are formed due to the presence of tight intercellular junctions between neighboring endothelial cells and interaction of the basolateral membrane of endothelial cells with the underlying basement membrane. Loss of barrier function or increased endothelial permeability is a hallmark feature of various diseases and clinical conditions such as severe trauma, sepsis, reperfusion injury, acute lung injury, acute respiratory distress syndrome (ARDS), and drug toxicity [1, 2]. Thus, studying the mechanisms of endothelial permeability and screening of potential drug compounds to prevent it requires a rapid and a sensitive method. Among the in vitro techniques available to assess barrier function, trans-well permeability assay, trans-epithelial electrical resistance (TEER) measurement, and electrical cell-substrate impedance sensing (ECIS) are in routine use by researchers

Binu Tharakan (ed.), Vascular Hyperpermeability: Methods and Protocols, Methods in Molecular Biology, vol. 2711, https://doi.org/10.1007/978-1-0716-3429-5_5, © Springer Science+Business Media, LLC, part of Springer Nature 2024

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worldwide. Unlike Transwell assays, TEER and ECIS provide rapid and label-free measurement of changes in epithelial/endothelial barrier function [3]. In this chapter we provide a detailed methodology for performing endothelial barrier function experiments for human umbilical vein endothelial cells (HUVECs) in response to thrombin with ECIS Model Zϴ. 1.1 Working Principle

ECIS is a multifrequency alternating current (AC) impedance measuring biosensor technology capable of measuring as well as modeling different cellular parameters in real time in a label-free manner. It was invented by Glaever and Keese and is commercially available from Applied BioPhysics, Inc. [4–6]. The ECIS system consists of a system controller, 16 or 96 well array station, and a computer (Fig. 1). This system can carry out through impedance spectroscopy; temporal monitoring of cellular behavior such as cell adhesion, migration, proliferation, and spreading; as well as barrier properties. Here the cells are grown in culture dishes carrying goldplated surfaces at the base of the wells (Fig. 2A). These gold surfaces are covered by thin insulating films, and gaps in these insulating films form parallel working electrodes which in turn are connected to a large in-plane counter electrode. Upon seeding, as the cells start spreading and growing over these electrodes, they act as insulators and impede the flow of current, thus creating a typical sigmoid curve (Fig. 2B). This curve has three phases: uncovered electrode (Z = Z0) showing 0 impedance value, change in impedance (Z = Z1) due to cell seeding and subsequent establishment of

Fig. 1 The ECIS Zθ system: (A) ECIS Zθ controller setup. (B) 16- and 96-well array station. Images reproduced with permission from the Applied BioPhysics website. (https://www.biophysics.com/public/pdf/ ECISProductGuide.pdf)

Determination of Endothelial Barrier Resistance by ECIS System

A

B

49

Cell manipulation

Cell Seeding

C

Medium Rb

Rb Cm

Cm

α Electrode

Fig. 2 Working principle of ECIS system: (A) A 96-well plate and an enlarged image of one of its well showing internal circuit diagram. (Taken from the website of Applied Bio Physics with permission). (B) Schematic representation of a typical sigmoid curve obtained by ECIS system. (Taken from the website of Applied Biophysics with permission). (C) Schematic representation of the working principle of ECIS measurement

confluent monolayer, and finally change in impedance (Z = Z2) due to experimental treatment of cell monolayer exhibiting barrier properties (Fig. 2B). The impedance thus generated is measured as per Ohm’s law, and as shown in Fig. 2C it is due to: 1. Capacitance contributed by the apical and basal membranes of cultured cells (Cm) which reflects the total area of electrode covered by them. 2. Resistance (Rb) provided by the paracellular spaces due to formation of intercellular tight junctions between neighboring cells. 3. Resistance (α) due to the strength of cell attachment with the basement matrix. Measurement of complex impedance in response to multifrequency AC and its breakdown into contributing factors of resistance (R) and capacitance (C) is a unique feature of ECIS, which is not applicable for other conventional models of TEER measurements. Hence, this technology allows for determination of relative contribution of paracellular junctions, strength of basal endothelial adhesion, and the capacitive behavior of cell membrane toward the overall impedance. The advanced ECIS Zθ Model allows for

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segregation of impedance into the contributing components of capacitance and resistance in real time based on mathematical modeling proposed by Glaever and Keese [4]. Application of weak AC allows for measuring impedance toward flow of electrons through and between adjacent cells (Fig. 2C). At low frequency (100–5000 Hz), majority of the AC flows between adjacent cells, thereby allowing measurement of paracellular barrier properties as resistance (R). At high frequencies (>104 Hz) the impedance is largely contributed by the capacitance (Cm) which allows measurement of cell adhesion, spreading, migration, and cell loss. In simpler terms, Rb allows assessment of tightness of intercellular spaces, α estimates changes in basal adhesion and cell size, and Cm measures cellular capacitance [4, 7]. For successful estimation of resistance due to paracellular barrier in the ECIS system, the following conditions should be satisfied: 1. Electrodes should be thoroughly covered by a tight cell monolayer. 2. Cells should be of uniform morphology and size and they should all be at similar height above the electrode. 3. Current density should be constant and should flow through space between neighboring cells at low frequency. 1.2 Prerequisite Considerations and Optimization

Given that endothelial cells across vasculature exhibit phenotypic heterogeneity, it is important to have a fair idea about the time taken by a particular type of endothelial cells to form a tight monolayer and to develop a functional resistant barrier. Even for a given cell type, it is not necessary that the time taken to form a monolayer will coincide with the establishment of a resistant barrier. For instance, although a monolayer is established within 20 h of seeding at a seeding density of 20,000 cells/well for human cerebral microvascular endothelial cells, the functional barrier appears 40–50 h after seeding and it remains stable for the next 50 h [8]. Hence, before conducting exploratory experiments for resistance measurements in a single frequency mode, an investigator should acquire impedance data at multi-frequency and model it to determine Rb, Cm, and α for cell-free and cell-covered electrodes. Modeling in this manner allows determination of time taken for appearance and establishment of a stable barrier (Rb). It also provides information about the duration of stability of the barrier to conduct experiments. As a guiding principle in the ECIS Zθ system, resistance measurement can be performed for most of the cell lines at a single frequency of 4000 Hz, while capacitance can be measured at higher frequency of 10–60 KHz.

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Materials

2.1 Instrument Requirement

1. ECIS Model Zθ system controller (Applied BioPhysics Inc., Troy, NY), as shown in Fig. 1A. 2. 96-well array station (Applied BioPhysics Inc., Troy, NY), as shown in Fig. 1B. 3. 96W20idfPET array plate (Applied BioPhysics Inc., Troy, NY) as shown in Fig. 2A. 4. Integrated software for 96-well array station, Version-1.2.254 that runs on Windows. 5. Cell culture CO2 incubator with HEPA filtration system and humidity controller. It is preferable to have water-jacketed incubators as they maintain desirable temperatures for longer duration. 6. A Class II Vertical-Type Laminar Flow Hood with HEPA filtered air flow. 7. A phase contrast cum immunofluorescence microscope. 8. Centrifuge with swing-out bucket rotor. 9. Automatic cell counter or hemocytometer.

2.2 Buffers and Solutions

1. Prepare all solutions in autoclaved ultrapure water (collected from Milli-Q water purification system with a sensitivity of 18 MΩ.cm at 25°C). Do not prepare reagents in phosphate buffer saline (PBS) as it can interfere with adsorption of proteins onto the array electrodes. 2. Before using any solution for experiment, make sure they are prewarmed at 37°C in water bath for at least 30 min. Adding cold solution will lead to large changes in resistance values. 3. 1 mg/mL gelatin (Sigma, St. Louis, MO) in HEPES buffer. Alternatively, the base of wells can be coated with fibronectin solution (dissolve 1 mg fibronectin in 32 mL of sterile distilled water to get a working concentration of approximately 30 μg/ mL). 4. 10 mM L-cysteine (Sigma, St. Louis, MO) in Milli-Q water. 5. 1XHEPES buffer free of Ca2+ and Mg2+ions, pH 7.4 from Thermo Fisher Scientific, Waltham, MA, USA. 6. Human umbilical vein endothelial cells (HUVECs) from Invitrogen, Carlsbad, CA, USA. Primary HUVECs can be used up to passage number 5–6. 7. EGM™ Endothelial Cell Growth Medium Bullet kit with fetal bovine serum from LONZA (CC-3124).

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Methods Instrument Setup

For ECIS array holder, choose a dedicated CO2 incubator with 5% CO2 levels, temperature of 37°C, and humidity levels of about 95%. Water tray filled with sterile distilled water usually contributes to 90–95% of humidity through passive evaporation. The array holder must be placed in CO2 incubator for 1 h and must be connected to the ECIS device along with a personal computer (PC) prior to start of an experiment. For endothelial permeability experiments, it is always better to use arrays with multiple electrodes. Here we describe experimental protocol for 96W20idfPET plate, as it has maximum electrode area and thus measures the impedance for maximum number of cells allowing minimum variability. Each well of this array contains electrodes in interdigitated configuration providing a total electrode area of 3.9 mm2 which can measure around 8000 cells at a time (Fig. 2A) for a confluent monolayer. Please go through the ECIS website for more information on availability of various types of culture arrays (https://www. biophysics.com/public/pdf/ECISProductGuide.pdf).

3.2 Calibration with Test Array (To Locate Respective Buttons on the Software, See Fig.3)

1. Open the ECIS software and press SETUP , to establish the connection between the array holder and the computer.

3.3 Coating of Array Wells

1. Coat the plate with 100 μL of 1% gelatin solution or fibronectin (32 μg/mL) in each well and incubate the array plate for 30 min at 37°C in CO2 incubator.

3.1

2. Place the test plate array into the array holder and ensure connection of the pins. Do not tighten the knob too much. 3. Select “RC test” from the type of array and press CHECK button and wait for the instrument to complete the run. On completion, the instrument will display the known impedance, resistance, and capacitance values for 96 wells mentioned in the user manual.

2. Remove the gelatin or fibronectin solution and add 100 μL of complete cell culture medium in each well. 3. Place the cell plate array into the array holder and make sure to match the connection of the pins. Do not tighten the knob too much. If the user does not wish to use entire plate, then all stabilization and further processing should be done for the selected wells based on the experimental plan. It is always better to have a 96-well plate print and label your treatment plan. 4. Open the ECIS software and press SETUP , to establish the connection between the plate array and the computer. Follow this with selection of the type of array as 96W20idfPET and press CHECK to confirm the connectivity of the instrument to all the wells. In the software panel, under “well

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Fig. 3 ECIS software window showing SET-UP menu. Red arrow displays the positions of key buttons. Taken from Applied BioPhysics instruction manual with permission

configuration,” green well indicates good connection, while red well indicates no connection (Fig. 3). 5. After all wells light up green, check the resistance/impedance values for all the wells. There should be minimum variation across the wells for the entire plate. If there is a lot of variation, you need to stabilize the electrodes by using the stabilization function of the setup. For this ECIS will automatically apply a high current to clean the electrodes. Press STABILIZE function and wait until the process is completed. Press CHECK again to see the stabilization of electrodes. Alternatively, one can perform L-cysteine treatment as listed in Notes section. 3.4 Seeding of the Cells

Perform all manipulations in a laminar flow cabinet under sterile conditions. 1. For performing barrier function experiments, employ HUVECs within passage 5–6 which were maintained in a gelatin or fibronectin coated T-25 flasks and were grown to confluence in growth medium with 10% FBS. Confluent monolayer should be washed with 1× HEPES Ca2+ and Mg2+ ions free buffer. Cells should be trypsinized as routinely done in

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cell culture during passaging of cells to obtain a homogenous cell suspension in a 15 mL falcon tube. Pellet the cells by centrifugation at 1000 rpm (130 × g) and resuspend the cells in 1 mL endothelial growth medium. Count the cells with a hemocytometer or an automatic cell counter (see Notes 1 and 2). 2. For the permeability assay, endothelial cells should be seeded at an approximate cell density of 30,000–40,000 cells/well. Set aside few control wells, where no cells are added and only 150 μL of growth medium is added. It is always better to prepare cells in such a way that adding 150 μL to each well will result in a desirable seeding density. For instance, prepare 2.5 × 105 cell/mL cell suspension. Adding 150 μL to each well from this cell suspension will result in approximately 37,000 cells/well. Before seeding the cells onto the wells of the array plate, make sure that the cells are uniformly suspended and are not clumped together (see Notes 3 and 4). 3. Select the wells to be measured in the well layout section. 4. Press CHECK button again. Once all wells light up green, proceed further otherwise check all connections. 5. Following addition of 150 μL of cell suspension, the final volume in each well will be 250 μL. It is preferred to conduct experimental treatment in duplicate wells. This is regarded as technical replicates. For confirming reproducibility of data, experiments should be performed multiple times with different batches of HUVECs, every time in technical duplicates. 6. Select Single Frequency Time (SFT) and choose the measurement frequency of 4000 Hz from the drop-down menu (Fig. 3). There is no need to adjust for time settings as the instrument will adjust to seconds based on number of wells and number of frequencies. If the experimenter wishes to customize these settings, these can be edited by the user in the commands. Low frequencies (4000 Hz) are best for endothelial barrier studies as majority of the current passes through the paracellular route [8]. If the investigator is keen to determine Rbas well as α, then impedance measurements should be performed in multiple frequency mode (MFT), and the acquired impedance data should be modeled and analyzed as described in the ECIS instruction manual (see Notes 5 and 6). 7. Press START and follow the instructions and save the experiment details in a file along with the experiment date on the computer. Incubate cells for the next 48 h and follow their adhesion curve. It takes anywhere between 30 and 48 h for cells (HUVECs) to form a stable barrier (see Note 7). 8. Before performing experiments with agonist treatment, it is necessary to determine for how long the barrier remains stable for a given cell type. Usually, the barriers remain stable for up to

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50 h after reaching plateau and this time window is sufficient to perform experimental manipulations. For HUVECs, it usually takes 24 h to reach plateau for the growth curve if the initial cell seeding density is 30,000–40,000 cells/well. However, it is always better to wait for additional 24 h (total 48 h) to see optimal barrier formation through formation of tight junctions. For other cells, it is advisable to perform a cell titration step to determine best seeding density for reaching a plateau curve (see Notes 8 and 9). 3.5 Medium Change and Thrombin Treatment

1. After formation of a tight barrier (plateau) for HUVECs (Fig. 2B), Press PAUSE to remove the array from the holder. 2. Place the array plate into the laminar flow hood and remove the culture medium and add 200 μL of prewarmed cell culture medium containing 1% serum. This step is done to serum starve the cells prior to subjecting them to an agonist treatment (in this case thrombin). 3. Place array back in the holder and press RESUME to continue data acquisition. Wait for the stabilization of resistance/ impedance values. Wait for a minimum of 3 h to serum starve the cells and to stabilize the changes on the electrodes. Press PAUSE and remove the array. Serum starvation is not a necessary step. If the plan is to carry out measurement for 2–3 days, it is desirable to replace with fresh endothelial growth medium every 2 days. It should be noted that following medium change, it usually takes 30 min for barrier readings to stabilize. 4. Now, add the agonist (thrombin) from the stock solution directly to the well. Here, we added thrombin at a final working concentration of 30 ng/mL. Note down the time in your notebook, or this can be marked in the curve as a comment through software. The vertical green line marked in Fig. 4 indicates medium change and treatment with thrombin. 5. Press RESUME to continue data acquisition. This experimental step can be marked by clicking the MARK button on the software and entering the details. The duration of data acquisition depends on the type of agonist that is being tested. Since thrombin causes rapid changes in endothelial permeability which last for about an hour, data acquisition can be stopped upon recovery of the barrier function. However, in case of TNFα, where there is a gradual loss of VE cadherin by endocytosis, it may take almost 12–18 h to see an appreciable change in permeability [9, 10]. Thus, it is important to have some idea about the permeability inducing effects of the agonist that is being studied (see Notes 10 and 11).

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Data Analysis

Through software: 1. To analyze ECIS results, press ANALYZE and select the resistance R from the top of the menu as shown in Fig. 3. 2. Choose exact timepoint to see your treatment effects by using the “Offset and Range” sliders beneath the graph, limit the starting time and the length of time to be analyzed. Here for example, we are showing thrombin experiment (Fig. 4A). After limiting the time, you will see clear graph for thrombin effects on endothelial permeability (Fig. 4B). 3. Now, press “Normalize” function, or press the n/n0 icon to normalize the curve (this divides all the obtained resistance value by the resistance values obtained at the time listed in the “zero time” box. This refers to the time point at which thrombin was added. In the dialog box of “zero time,” add the time at which thrombin was added (e.g., 47.2 h for this experiment). This will default set the time of thrombin addition as time “zero” (Fig. 4C). To accommodate for changes among

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different wells, resistance values need to be normalized. Normalization is done for a specific time point, where all values are set as 1.0 by the software. For instance, for the testing of adhesion of cells on various matrices, normalization should be done at time 0, where cells were added. Then we will be able to see the clear difference on their adhesion ability on various matrix proteins. In the current case of thrombin treatment, normalization should be done at time of thrombin addition (Fig. 4). Thus, this brings all changes before the addition of thrombin to 1. Hence, the user can now see the clear effect of thrombin on endothelial permeability. 4. Other optional steps: By pressing “Running Average,” data will get smooth by entering specified number of data points in the drop-down menu. 5. By pressing “Bin Running Average,” data points can be reduced by displaying every n-th point. 6. Press “Standard Deviation,” to show standard deviation bars. Manual Data Analysis Another option to analyze your results is to export all the results in MS Excel and save it. In the exported Excel sheet, all individual impedance, resistance, and capacitance values will be depicted. Choose resistance values along with all-time points and plot a curve. A typical curve involves a cell attachment phase, cell spreading/cell growth phase, and a stabilized barrier phase (Fig. 5A). Then treatment timepoint is selected (Fig. 5B) and resistance values are normalized manually (Fig. 5B, C). All resistance values are normalized by dividing them with resistance value observed at the time of thrombin addition. These tracings are plotted as normalized resistance with respect to experimental control (i.e., no thrombin treatment). The curve can be further resolved to exact time point, where thrombin response started and ended (Fig. 5D). Furthermore, maximum decrease in normalized resistance can be plotted as a bar graph from treated and untreated well (Fig. 5E). Here the lowest resistance value is chosen for the preparation of bar graph from both the groups by Graph Pad Prism. 3.7 Data Modeling for Determination of Rb and α

1. The ECIS Zθ software can calculate time-dependent changes in the model parameters (Rb and α) as discussed earlier. To accomplish this, it is necessary to keep one well cell-free and record the time course data using multiple frequencies (MFT) instead of SFT. 2. The cell-free culture medium-filled well is selected as a reference well. Press FIND to automatically select the reference value. The instrument will automatically find the cell-free well. It is also possible to select “cell free time point” in a well manually by using the “frequency scan modeling” available in the ANALYZE section.

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3. If a cell-free reference is not available, one can import reference value from another experiment, provided they were conducted on the same type of array by using the same culture medium. To find the cell-free data from another experiment, open that experiment file. Then select the cell-free well by cursor or by pressing find. Now, click Set and return to the dataset, which needs to be modeled and click Get. 4. For data modeling, after finishing the data recording, follow these steps. Choose the time frame by using the Offset and Range sliders during which the modeling must be done. 5. Press MODEL to start the calculations. This will open another window (Fig. 6A, B). Select cell-free reference: (CFR) media-only well and cross check the well number under “current selection.” Then press final model button. Time taken to perform this action depends upon the size of the acquired dataset. At the end, from the original curve (Fig. 6C) instrument will generate the curves for Rb and α (Fig. 6D) and should display the corresponding values. High Rb (Ω.cm2) indicates low permeability toward current flow.

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Notes 1. For measurement of barrier properties for endothelial cells, it is important to bear in mind that throughout the vasculature, endothelial cells exhibit heterogeneity. Hence, for a given endothelial cell source, it is necessary to have some idea about the relative contribution of paracellular junctions versus basal adhesion toward barrier properties. For instance, in lymphatic endothelial cells the contribution of basal cell adhesion to ECM is more compared to that from the intercellular junctions. In the blood-brain barrier (BBB), intercellular tight junctions play a crucial role in determining barrier property. It is also worth noting that for permeability studies, cell lines do not express barrier properties at par with in vivo conditions and thus primary cells are preferred. 2. It is important for seeded cells to reach confluence and establish a stable barrier resistance before testing for effects of

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agonists or experimental treatments. Change in impedance for sub-confluent cultures is largely related to cell proliferation. 3. Physiologically, the apical membrane of brain endothelial cells is exposed to serum compared to the abluminal side and thus ECIS is a preferred system to study BBB. Proximity of cells to the electrodes when grown on ECIS arrays provides high sensitivity of measurement for intercellular barrier. The basolateral fluid compartment is negligible in ECIS system, and thus it cannot be used for performing transcellular transport experiments. 4. Primary endothelial cells are usually cultured in enriched cell culture medium supplemented with growth factors, serum, and other media supplements such as hydrocortisone, heparin, ascorbic acid, etc. Even subtle differences in culture medium can influence the barrier properties of confluent endothelial monolayers. For instance, serum, growth factors, cAMP, glucocorticoids, and hydrocortisone are known to strengthen barrier properties [11–13]. A shift from 10% FBS to 2% FBS can cause a 20% dip in Rb, and thus upon changing the media, one should wait for the impedance to stabilize before starting agonist treatments. Other agents which may modulate the barrier are D-mannitol and DMSO. 5. Electrode stabilization by L-cysteine treatment: This process is known to improve the stability of electrode by direct binding of L-cysteine to gold surface through thiol groups [14]. For this, add 100 μL of 10 mM L-cysteine solution to each well and incubate the plate for 15 min at room temperature. Wash three times with 200 μL/well Milli-Q water. 6. As mentioned earlier, it is desirable to record resistance offered by endothelial monolayer in SFT mode which also provides higher temporal resolution. If it is not known as to which frequency is ideal for conducting SFT measurements for the type of endothelial cells that are under investigation, one can perform frequency scan. In the resistance versus frequency log plot, the frequency at which the difference between the cellfree versus cell-covered electrode is maximum, that frequency can be regarded as the optimal frequency at which the cells impede current effectively to conduct barrier function measurements. For endothelial cells, this frequency is 4000 Hz. 7. A good resistance/impedance range for formation of strong barrier is 400–1000 Ω. The duration for which this barrier will remain stable however varies from cell type to cell type. Thus, it is necessary to have an idea about the following for a given cell type before executing exploratory experiments: (a) Average time taken by an endothelial cell type to form barrier.

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(b) Temporal stability of barrier to provide experimental time window to perform manipulations and treatments. (c) Culture conditions to achieve a strong and stable barrier for a given endothelial cell type. 8. It is important to revalidate data obtained from ECIS on changes in barrier property by performing immunofluorescence assessment of junctional proteins such as VE-cadherin (CD144) and zona occludens-1 (ZO-1) in parallel. 9. ECIS Applied BioPhysics, Inc. provides gold electrode culture arrays in different configurations including 8-well array, 16-well arrays, 96-well microtiter plate arrays, or specialized flow arrays from Ibidi GmbH. In 8W10E array there are ten circular electrodes connected in parallel in which for a confluent cell monolayer, the electrodes can measure 500–1000 cells for different cellular functions. In 96W20idfPET, each well has an electrode coverage area of 3.9 mm 2 measuring 4000–8000 cells at a time (Fig. 2A). This large collection of small electrodes thus allows for improved sampling with minimal variation. 10. In the ECIS Zθ system during single frequency (SFT) run, each well will be recorded at a specified frequency, while during the multiple frequency (MFT) run, the Zθ program measures each well at seven different predefined frequencies for 96-well arrays. For each of these modes, the minimal time interval between two successive measurements and the overall duration of recording can be specified in the software. The acquisition rate for a 96-well array in ECIS Zθ for SFT is 0.25 s/well and 7.5 s/well for MFT. The rapid time collect (RCT) mode is employed when rapid data acquisition is needed for agonists which elicit very quick changes. These measurements for RCT are performed in single well at a time. 11. The frequency range available for experimentation in ECIS Zθ is from 25 Hz to 100 KHz. References 1. Baluna R, Vitetta ES (1997) Vascular leak syndrome: a side effect of immunotherapy. Immunopharmacology 37(2–3):117–132. https:// doi.org/10.1016/s0162-3109(97)00041-6 2. Duan CY, Zhang J, Wu HL, Li T, Liu LM (2017) Regulatory mechanisms, prophylaxis and treatment of vascular leakage following severe trauma and shock. Mil Med Res 4:11. https://doi.org/10.1186/s40779-0170117-6 3. Bischoff I, Hornburger MC, Mayer BA, Beyerle A, Wegener J, Furst R (2016) Pitfalls in assessing microvascular endothelial barrier

function: impedance-based devices versus the classic macromolecular tracer assay. Sci Rep 6: 23671. https://doi.org/10.1038/srep23671 4. Giaever I, Keese CR (1991) Micromotion of mammalian cells measured electrically. Proc Natl Acad Sci USA 88(17):7896–7900. https://doi.org/10.1073/pnas.88.17.7896 5. Giaever I, Keese CR (1984) Monitoring fibroblast behavior in tissue culture with an applied electric field. Proc Natl Acad Sci USA 81(12): 3761–3764. https://doi.org/10.1073/pnas. 81.12.3761

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6. Giaever I, Keese CR (1993) A morphological biosensor for mammalian cells. Nature 366(6455):591–592. https://doi.org/10. 1038/366591a0 7. Benson K, Cramer S, Galla HJ (2013) Impedance-based cell monitoring: barrier properties and beyond. Fluids Barriers CNS 10(1):5. https://doi.org/10.1186/20458118-10-5 8. 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). https://doi.org/10.3390/ bios8030064 9. Adam AP, Lowery AM, Martino N, Alsaffar H, Vincent PA (2016) Src family kinases modulate the loss of endothelial barrier function in response to TNF-alpha: crosstalk with p38 signaling. PLoS One 11(9):e0161975. https:// doi.org/10.1371/journal.pone.0161975 10. Petrache I, Birukova A, Ramirez SI, Garcia JG, Verin AD (2003) The role of the microtubules in tumor necrosis factor-alpha-induced endothelial cell permeability. Am J Respir Cell Mol

Biol 28(5):574–581. https://doi.org/10. 1165/rcmb.2002-0075OC 11. Hoheisel D, Nitz T, Franke H, Wegener J, Hakvoort A, Tilling T, Galla HJ (1998) Hydrocortisone reinforces the blood-brain properties in a serum free cell culture system. Biochem Biophys Res Commun 247(2): 312–315 12. Kroll S, El-Gindi J, Thanabalasundaram G, Panpumthong P, Schrot S, Hartmann C, Galla HJ (2009) Control of the blood-brain barrier by glucocorticoids and the cells of the neurovascular unit. Ann NY Acad Sci 1165: 228–239. https://doi.org/10.1111/j. 1749-6632.2009.04040.x 13. Weidenfeller C, Schrot S, Zozulya A, Galla HJ (2005) Murine brain capillary endothelial cells exhibit improved barrier properties under the influence of hydrocortisone. Brain Res 1053(1–2):162–174. https://doi.org/10. 1016/j.brainres.2005.06.049 14. Tengvall P, Lestelius M, Liedberg B, Lundstrom I (1992) Plasma-protein and antisera interactions with L-cysteine and 3-mercaptoproprionic acid monolayers on gold surfaces. Langmuir 8(5):1236–1238. https://doi.org/10.1021/la00041a001

Chapter 6 Time-Lapse Observation of Cell Dynamics During Angiogenesis Using the Rat Mesentery Culture Model Arinola O. Lampejo, Nicholas A. Hodges, Maximillian Rozenblum, and Walter L. Murfee Abstract The ability to track cells and their interactions with other cells during physiological processes offers a powerful tool for scientific discovery. An ex vivo model that enables real-time investigation of cell migration during angiogenesis in adult microvascular networks would enable observation of endothelial cell dynamics during capillary sprouting. Angiogenesis is defined as the growth of new blood vessels from existing ones and involves multiple cell types including endothelial cells, pericytes, and interstitial cells. The incorporation of these cell types in a physiologically relevant environment, however, represents a challenge for biomimetic model development. Recently, our laboratory has developed the rat mesentery culture model, which enables investigation of angiogenesis in an intact tissue. The objective of this chapter is to detail a protocol for tracking cellular dynamics during angiogenesis using the rat mesentery tissue culture model. The method involves harvesting mesentery tissues from adult SD-EGFP rats, culturing them in MEM + 10% fetal bovine serum, and imaging network regions over the time course of angiogenesis. In example applications, time-lapse comparison of microvascular networks in cultured tissues confirmed dramatic increases in GFP-positive capillary sprouting and GFP-positive segment density. Additionally, tracking of individual capillary sprout extensions revealed their ability to “jump” by disconnecting from one vessel segment and reconnecting to another segment in the network. GFP-positive sprouts were also capable of undergoing subsequent regression. The representative results support the use of the rat mesentery culture model for identifying and tracking cellular dynamics during angiogenesis in intact microvascular networks. Key words Microcirculation, Angiogenesis, Time-lapse imaging, Microvascular remodeling, Tissue culture, Mesentery, Capillary sprouting

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Introduction Understanding the cellular dynamics involved in microvascular growth is dependent on the ability to watch cells during angiogenesis, defined as the growth of new capillaries from existing vessels. Essential to this process are endothelial cells and their ability to sprout or extend from one vessel to another. Endothelial cell sprouting dynamics are thus critical not only in angiogenesis but also in multiple disease states such as cancer metastasis, ischemia,

Binu Tharakan (ed.), Vascular Hyperpermeability: Methods and Protocols, Methods in Molecular Biology, vol. 2711, https://doi.org/10.1007/978-1-0716-3429-5_6, © Springer Science+Business Media, LLC, part of Springer Nature 2024

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and a variety of inflammatory disorders [1]. The behavior of endothelial cells in the context of physiological or pathological angiogenesis is influenced by other cell types such as vascular pericytes, immune cells, and local interstitial cells [2]. Furthermore, endothelial cell phenotypes and functions differ depending on their location within a network [3]. These cell-cell interactions and dependence on local microenvironments motivate the need to not just watch endothelial cells but to watch endothelial cell dynamics in an intact microvascular network. Current in vitro, ex vivo, and in vivo methodological models have provided insights into endothelial cell dynamics during angiogenesis. Notably, transgenic zebrafish models allow for noninvasive, prolonged, in vivo, time-lapse imaging of vasculature at any stage of development [4–6]. Further, multiphoton imaging approaches have also been applied to observe vessel growth in mouse pial vasculatures [7, 8]. In vitro angiogenic models include three-dimensional tubule formation assays [9] and microfluidic platforms [10–12]. Examples of ex vivo models include culturing mouse retinal tissues [13], the chorioallantoic membrane assay [14], and the aortic ring assay [15]. Each angiogenesis assay is highlighted by strengths and limitations. Incorporating the complexity of a real microvascular network with the ability for time-lapse imaging remains a common challenge in biomimetic model development. As an alternative approach to meet this challenge, our laboratory has introduced the rat mesentery culture model (Fig. 1) [2, 16]. The rat mesentery is relatively easy to harvest, and the thinness of the rat mesentery (20–40 μm) allows for imaging intact microvascular networks [18]. This model also contains a variety of other cell types including pericytes, smooth muscle cells, and interstitial cells [2], all of which can be studied to determine what roles they play in angiogenic growth. Characteristics of the rat mesentery culture model include (1) the maintenance of endothelial cell, pericyte, and smooth muscle cell viability [2], (2) the maintenance of immune cell, interstitial cell, and lymphatic endothelial cell viability [16], (3) the maintenance of blood and lymphatic networks as well as the ability to induce angiogenesis [17], and (4) the ability to induce lymphangiogenesis and track endothelial cell segments over time [16]. The objective of this chapter is to detail a protocol for tracking cellular dynamics during angiogenesis using harvested mesentery tissues from GFP-transgenic rats. Our representative results show examples of capillary sprouting across the hierarchy of microvascular networks and highlight the value of observing angiogenesis to make discoveries of new endothelial cell dynamics.

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Fig. 1 Overview of the rat mesentery culture model. Mesentery tissues are excised, rinsed, inserted into well plates, and sandwiched between the commercially available membrane insert and the well bottom. The use of tissues harvested from GFP-transgenic rats enables the microscopic observation of intact microvascular networks. Scale bar = 250 μm

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Surgical Tools

1. Hair clippers. 2. Scalpel handle. 3. Sterile surgical scalpel blade. 4. Surgical scissors. 5. Microdissecting scissors. 6. Microdissecting forceps. 7. 60 mm Petri dish. 8. Sterile, tissue-culture treated six-well plate.

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1. Assemble the CellCrown inserts (see Notes 1–2). 2. Sterilize all surgical tools and materials in autoclave pouches (see Notes 3–8). 3. In 50 mL sterile conical tubes, prepare 30 mL of sterile saline. 4. In two petri dishes, prepare 10 mL of DPBS and 10 mL of MEM supplemented with 1% Pen-Strep. 5. Incubate all solutions in a water bath set to 37 °C. 6. Prepare the culture media (4 mL for every tissue collected) and store at 4 °C. 7. Prepare the surgical bench by wiping the station with 70% ethanol. 8. Spread the sterile absorbent pad onto the surgical station and place the plexiglass plate on top of the pad. 9. Set up the electric heating pad near the workspace, and place sterile drapes onto the pad and on the other side of the station. 10. Place the sterile surgical tools on the other sterile drape.

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1. Anesthetize the rat (see Note 9). 2. Wait for approximately 5 min and confirm that the rat is under anesthesia by pinching between the animal’s toes and checking for an auditory response. These tests should not cause any reflexes. 3. Shave the abdominal area with clippers to remove the majority of the hair. Spread Nair over the shaved area, and let sit for 2 min to ensure complete removal of hair. 4. Wipe the Nair off with gauze. Once most of the Nair has been removed, wipe the rat using 70% isopropanol and povidoneiodine solution twice (see Note 10). 5. Transfer the rat to the surgical station, laying it supine on the plexiglass. 6. Turn on the heating pad and place the two petri dishes onto the pad. Place the warmed saline by the surgical station.

3.3 Mesenteric Tissue Exteriorization

1. Below the sternum, use a scalpel blade to make a longitudinal incision through the skin and abdominal wall. 2. Once the abdominal cavity is opened, use surgical scissors to cut a 2-inch incision through the rest of the tissue. 3. Place the precut drape over the abdominal section so that the opening aligns with the incision. 4. Using cotton-tip applicators locate the cecum and pull it to the opening. 5. Place the sterilized surgical stage overtop the opening and wet it with saline. 6. Pull out the cecum and place it on the stage. Use the cotton-tip applicators to pull the rest of the small intestine while also identifying vascularized mesenteric windows. Avoid touching the mesenteric windows. 7. Euthanize the rat (see Note 9). 8. Ensure the rat has no pulse before continuing with the harvesting process.

3.4 Harvesting Mesentery Tissue

1. Locate a desired mesenteric window and spread it out fully, making sure not to touch the window in the process. 2. At the top of the mesenteric window, use the microforceps to grip the fat and make an incision with the microscissors just behind microforceps. 3. Use the microscissors to cut out the window (see Notes 11– 12). 4. Rinse the mesenteric window by immersing it in the petri dish of warmed DPBS.

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5. Transfer the mesenteric window to the petri dish containing media. 6. Repeat steps 1–4 until all desired mesenteric windows have been harvested. 3.5 Mounting the Tissues for Culture

1. In a biosafety cabinet, take one CellCrown out and tighten the membrane by pressing on the edges. 2. With a sterile microforcep, take out one of the mesenteric windows and gently blot it on the drape before placing it on the membrane in order to remove excess fluid. 3. Spread out the tissue over the membrane by using the straight microforceps to nudge the window apart while the flat part of the angled forceps holds the opposite edge of the peripheral fat pad. Spread out the window so the tissue is fully exposed and flattened (Fig. 1). 4. Add 1 mL of media to the well and then carefully place the insert in the well by inverting it and gently sandwiching the tissue between the insert and the bottom of the well. 5. Add an additional 3 mL media to the sides and center of the well. 6. Repeat steps 1–5 for all tissues. 7. Transfer the plate to an incubator under standard culturing conditions (see Note 13).

3.6 Imaging and Observation of Microvascular Networks

1. Since tissues are harvested from a GFP-transgenic rat strain and the tissues are approximately 20–40 μm thin [18], the microvascular networks can be imaged with a standard epifluorescent microscope as you might a monolayer of fluorescently tagged cells during typical cell culture (see Note 14; Fig. 2). 2. Depending on the imaging intervals, time-lapse imaging can be done with or without a temperature and CO2 control incubation chamber. 3. Images can be taken as often as every hour for up to several days (see Note 15; Figs. 2, 3, and 4).

3.7 Post-imaging Immunohistochemistry

1. Spread tissues onto microscope slides, cutting away the fat border of each tissue. 2. Fix the tissues in 100% methanol for 30 min at -20 °C (see Note 16). 3. Perform three 10-min washes with PBS 1% saponin. 4. Tap dry each slide. 5. Using a Kimwipe or sterile cotton-tip applicator, carefully blot dry the slide area around the tissue. 6. Using a hydrophobic marker, circumscribe each tissue.

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Fig. 2 Time-lapse imaging of microvascular network growth. GFP expression by cell populations in cultured rat mesentery tissue enables time-lapse imaging of adult microvascular networks over the time course of angiogenesis. Angiogenesis due to serum stimulation is characterized by a dramatic increase in vessel density (a–f). Scale bars = 250 μm

Fig. 3 Time-lapse imaging of endothelial cell sprouting. Individual capillary segments can be identified as “jumping” from one vessel to another. (a) Representative images of the tissue were taken at Day 0 of culture. (b, c) The formation of sprouts and extension of sprouts can be seen on both of the two central vessels. (d) Vessel begins connecting with the other existing capillary vessels or sprouts. (e, f) Both sprouts detach from the vessels of origin at different timepoints and after different initial sprouting mechanisms. Solid arrows identify sprouts of interest and outlined arrows identify a disconnection site from the original vessel. Asterisks identify a connection site between a sprout and a nearby vessel or opposing sprout. Scale bars = 50 μm

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Fig. 4 Tracking network/sprout extension and retraction. (a) Representative image showing two sprouts that have formed a small, branched intersection. (b) The sprouts subsequently diverge. (c) Increased network growth then occurs before the sprouts (d) re-converge, this time forming a more stable branched intersection. Solid arrows identify timepoints where the sprouts are connected and outlined arrows identify timepoints where the sprouts are disconnected. Scale bars = 100 μm

7. Pipet primary antibody solution onto each tissue (approximately 100 μL) and incubate in the dark at room temperature for 1 h (see Note 17). 8. Turn the slide on its side and tap off the antibody solution. 9. Using a dropper to completely cover the tissues perform three 10-minute washes with PBS 1% saponin. 10. Add 100 μL of secondary antibody solution onto each tissue and incubate in the dark at room temperature for 1 h (see Note 18). 11. Turn the slide on its side and tap off the antibody solution. 12. Using a dropper to completely cover the tissues, perform three 10-min washes with PBS 1% saponin. 13. Remove the hydrophobic pen markings and use a disposable pipette to deposit a few drops of PBS-glycerol solution with a 1:1 ratio onto each tissue. 14. Use a glass slide cover to then seal the tissues, minimizing air bubbles.

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Fig. 5 Comparison of GFP labeling and immunolabeling. (a) Representative image of a microvascular network taken at Day 0 of culture. (b) Representative image of a microvascular network taken just before fixation. (c) Representative image of a microvascular network after immunolabeling with PECAM-positive endothelial cells shown in red and NG2-positive pericytes shown in green. Scale bars = 100 μm

15. Seal the slides with nail polish or alternative sealant. 16. After sufficient time to allow for the nail polish to dry, the tissues can be imaged (Fig. 5) or stored long term in a freezer (-20 °C). For short-term storage, slides with tissues can be stored in a refrigerator (4 °C). 3.8 Representative Observations

Microscopic observation of GFP cells identifies vessels across the hierarchy of intact microvascular networks (Fig. 1). Time-lapse imaging over the time course of 3 days documents increased capillary sprouting and vessel density (Figs. 2 and 3). GFP labeling also identified interstitial cell populations (Figs. 2 and 3) and lymphatic vessels (data not shown). The formation of blood capillary sprouts can be observed as early as day 1 when cultured with the 10% serum with more dramatic sprouting occurring by day 2. For this protocol description, 10% serum was used because it has been shown to be a robust stimulator of microvascular network growth in ex vivo culture [19–21]. The value of the time-lapse imaging protocol with GFP-transgenic labeled tissues, which enable iterative imaging over variable timepoints, is exemplified by the observation of novel dynamics. Focusing on individual capillary sprouts identifies characteristic endothelial extension and additional dynamics such as endothelial cell jumping, in which an endothelial cell disconnects from a capillary sprout before reconnecting to a different neighboring sprout (Fig. 3). Representative images show the formation of sprouts on two neighboring capillaries (Fig. 3b) which then extend (Fig. 3c) and connect with the neighboring vessel (Fig. 3d) and

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subsequently detach from the original vessel (Fig. 3e–f). Along with showing that this phenomenon exists, these images also highlight that endothelial cell jumping can occur in different ways. One sprout is shown extending and connecting to a different capillary before detaching from its original capillary. In the other example of sprout jumping shown in these images, two sprouts extend from different branches and connect, and then one of the sprouts detaches from its vessel of origin. Another endothelial cell dynamic involves the disconnection, extension, and subsequent reconnection of two capillary segments (Fig. 4). These observations question our full understanding of capillary sprout dynamics and highlight different modes for endothelial cells during angiogenesis beyond capillary sprouting. These observations demonstrate the simple application of the rat mesentery culture model for obtaining time-lapse images of angiogenic microvascular networks and the potential for making observations at both the network and cellular levels. Quantification of angiogenic metrics across timepoints can enable the comparison of specific growth factor effects or the evaluation of pro- or antiangiogenic drug testing. In addition, tissues can be immunohistochemically labeled to identify specific cell types post-imaging. For instance, Fig. 5 shows a GFP tissue after fixation and labeling for PECAM (endothelial cell marker) and NG2 (pericyte maker). Spatial mapping of GFP and PECAM suggests that the GFP labeling indeed identified endothelial cells. NG2 labeling supports the presence of pericytes along the vessels. Other components of the microvasculature, such as smooth muscle cells, lymphatics, interstitial cells, immune cells, and neurons can also be labeled for and imaged [2, 16].

4

Notes 1. CellCrown inserts with membrane filters can rip or wrinkle easily. It is recommended that two extra inserts are prepared in case some are damaged during preparation. 2. Recommended technique: Line the membrane up with the bottom insert of the CellCrown then, keeping fingers evenly spaced, press down the ring down with moderate pressure. Press and rotate fingers around to ensure a snug fit. If any membranes rip, reapply with a fresh membrane. 3. It is recommended that all surgical tools are prepared in three pouches and all mounting tools are prepared in two pouches. 4. The first surgical pouch should contain an absorbent pad.

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5. The second surgical pouch should contain all tools used during harvesting including one pair of microscissors, one pair of angled microforceps, one pair of surgical scissors, one scalpel handle, and three drapes. 6. The third surgical pouch should contain gauze sponges. 7. The first mounting pouch should contain all CellCrown inserts with membranes. 8. The second mounting pouch should contain tools used to mount the tissues to the CellCrowns including one drape, one straight microforceps, and one 90° microforceps. 9. All animal procedures should be approved by an institutional animal care and use committee (IACUC) or its equivalent. Animal experiments described in the current protocol were approved by University of Florida’s Institutional Animal and Care Use Committee. Adult rats were anesthetized via an intramuscular injection with ketamine (80 mg/kg body weight) and xylazine (8 mg/kg body weight). Rats were euthanized via intracardiac injection of Beuthanasia-D. 10. In order to reduce contamination, wipe in a circular motion from the center of the shaved area to the periphery. 11. Make sure to include a fat border around the mesenteric window to provide a cushion between the membrane insert and the well plate. 12. Cut evenly on both sides to minimize the likelihood of the window tearing. 13. Standard conditions: 5% CO2 at 37 °C, and 95% humidity. Media was changed every 2 days. 14. If transgenic rats are not being used, refer to previously detailed protocol which uses a BSI-lectin staining protocol to live image tissues [2]. 15. For a wider network view, imaging at 4× is recommended. For detailing smaller interaction, imaging at 10× is recommended. Using a microscope with a motorized stage is also recommended so that specific locations can be easily marked and imaged over the experimental time course. 16. GFP cell labeling disappears after methanol fixation, resulting in the ability to immunolabel the tissues with green fluorescent secondary antibodies. 17. The primary antibody solution detailed in this chapter consists of 5% normal goat serum (NGS), 1:200 biotinylated mouse anti-rat PECAM (BD Pharmingen; San Diego, CA) and 1:100 rabbit anti-mouse NG2 (MilliporeSigma; Burlington, MA) diluted in a solution of PBS containing 1% saponin and 2% bovine serum albumin (BSA).

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18. The secondary antibody solution detailed in this chapter consists of 5% NGS, 1:500 streptavidin secondary (Strep-CY2) (Jackson ImmunoResearch; West Grove, PA), 1:100 goat anti-rabbit secondary (GAR-CY3) (Jackson ImmunoResearch; West Grove, PA) diluted in a solution of PBS containing 1% saponin and 2% BSA.

Acknowledgments This work was supported by the National Institutes of Health under Award Number R01AG049821. The authors want to especially thank Dr. Christine Schmidt and her Biomimetic Materials & Neural Engineering Lab in the J. Crayton Pruitt Family Department of Biomedical Engineering at the University of Florida for their generous sharing of the GFP transgenic rats. A special additional thanks goes to Stacey L. Porvasnik and Dr. Nora Hlavac for their help with coordinating the collaboration. References 1. Potente M, Gerhardt H, Carmeliet P (2011) Basic and therapeutic aspects of angiogenesis. Cell 146(6):873–887. https://doi.org/10. 1016/j.cell.2011.08.039 2. Stapor PC, Azimi MS, Ahsan T, Murfee WL (2013) An angiogenesis model for investigating multicellular interactions across intact microvascular networks. Am J Physiol Heart Circ Physiol 304(2):H235. https://doi.org/ 10.1152/ajpheart.00552.2012 3. Carmeliet P (2005, December 15) Angiogenesis in life, disease and medicine. Nature. Nature Publishing Group. https://doi.org/10.1038/ nature04478 4. Lawson ND, Weinstein BM (2002) In vivo imaging of embryonic vascular development using transgenic zebrafish. Dev Biol 248(2): 307–318. https://doi.org/10.1006/dbio. 2002.0711 5. Cha YR, Weinstein BM (2007) Visualization and experimental analysis of blood vessel formation using transgenic zebrafish. Birth Defects Res C Embryo Today 81(4): 286–296. https://doi.org/10.1002/bdrc. 20103 6. Jung HM, Isogai S, Kamei M, Castranova D, Gore AV, Weinstein BM (2016) Imaging blood vessels and lymphatic vessels in the zebrafish. Methods Cell Biol 133:69–103. https://doi. org/10.1016/bs.mcb.2016.03.023 7. Shih AY, Driscoll JD, Drew PJ, Nishimura N, Schaffer CB, Kleinfeld D (2012, July 1)

Two-photon microscopy as a tool to study blood flow and neurovascular coupling in the rodent brain. J Cereb Blood Flow Metab. SAGE Publications Sage UK: London, England. https://doi.org/10.1038/jcbfm. 2011.196 8. Letourneur A, Chen V, Waterman G, Drew PJ (2014) A method for longitudinal, transcranial imaging of blood flow and remodeling of the cerebral vasculature in postnatal mice. Physiol Rep 2(12):e12238. https://doi.org/10. 14814/phy2.12238 9. Sanz-Nogue´s C, O’Brien T (2016) In vitro models for assessing therapeutic angiogenesis. Drug Discov Today. Elsevier Ltd. https://doi. org/10.1016/j.drudis.2016.05.016 10. Song JW, Munn LL (2011) Fluid forces control endothelial sprouting. Proc Natl Acad Sci U S A 108(37):15342–15347. https://doi. org/10.1073/pnas.1105316108 11. Moya ML, Hsu YH, Lee AP, Christopher CWH, George SC (2013) In vitro perfused human capillary networks. Tissue Eng Part C Methods 19(9):730–737. https://doi.org/10. 1089/ten.tec.2012.0430 12. van Duinen V, Zhu D, Ramakers C, van Zonneveld AJ, Vulto P, Hankemeier T (2019) Perfused 3D angiogenic sprouting in a highthroughput in vitro platform. Angiogenesis 22(1):157–165. https://doi.org/10.1007/ s10456-018-9647-0

Cell Dynamics During Angiogenesis 13. Gerhardt H, Golding M, Fruttiger M, Ruhrberg C, Lundkvist A, Abramsson A et al (2003) VEGF guides angiogenic sprouting utilizing endothelial tip cell filopodia. J Cell Biol 161(6):1163–1177. https://doi.org/10. 1083/jcb.200302047 14. Ribatti D, Nico B, Vacca A, Roncali L, Burri PH, Djonov V (2001) Chorioallantoic membrane capillary bed: A useful target for studying angiogenesis and anti-angiogenesis in vivo. Anat Rec 264(4):317–324. https://doi.org/ 10.1002/ar.10021 15. Nicosia RF (2009) The aortic ring model of angiogenesis: A quarter century of search and discovery. J Cell Mol Med. Wiley-Blackwell. https://doi.org/10.1111/j.1582-4934.2009. 00891.x 16. Azimi MS, Motherwell JM, Murfee WL (2017) An ex vivo method for time-lapse imaging of cultured rat mesenteric microvascular networks. J Vis Exp 2017(120):55183. https:// doi.org/10.3791/55183 17. Motherwell JM, Anderson CR, Murfee WL (2018) Endothelial cell phenotypes are maintained during angiogenesis in cultured

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microvascular networks. Sci Rep 8(1):5887. https://doi.org/10.1038/s41598-01824081-z 18. Norrby K (2006) In vivo models of angiogenesis. J Cell Mol Med 10(3):588–612. https:// doi.org/10.1111/j.1582-4934.2006. tb00423.x 19. Azimi MS, Myers L, Lacey M, Stewart SA, Shi Q, Katakam PV et al (2015) An ex vivo model for anti-angiogenic drug testing on intact microvascular networks. PLoS One 10(3):e0119227. https://doi.org/10.1371/ journal.pone.0119227 20. Motherwell JM, Azimi MS, Spicer K, Alves NG, Hodges NA, Breslin JW et al (2017) Evaluation of arteriolar smooth muscle cell function in an ex vivo microvascular network model. Sci Rep 7(1):1–12. https://doi.org/ 10.1038/s41598-017-02272-4 21. Azimi MS, Motherwell JM, Hodges NA, Rittenhouse GR, Majbour D, Porvasnik SL et al (2020) Lymphatic-to-blood vessel transition in adult microvascular networks: a discovery made possible by a top-down approach to biomimetic model development. Microcirculation 27(2). https://doi.org/10.1111/micc.12595

Chapter 7 Evaluation of Barrier Integrity Using a Two-Layered Microfluidic Device Mimicking the Blood-Brain Barrier Hossam Kadry and Luca Cucullo Abstract The blood-brain barrier (BBB) plays an essential role in maintaining the homeostasis of the brain microenvironment by controlling the influx and efflux of biological substances that are necessary to sustain the neuronal metabolic activity and functions. This barrier is established at the blood-brain interface of the brain microcapillaries by different cells. These include microvascular endothelial cells, astrocytes, and pericytes besides other components such as microglia, basal membrane, and neuronal cells forming together what is commonly referred to as the neurovascular unit; different in vivo and in vitro platforms are available to study the BBB where each system provides specific benefits and drawbacks. Recently, organon-a-chip platforms combine the elegance of microengineering technology with the complexity of biological systems to create near-ideal experimental models for various diseases and organs. These microfluidic devices with micron-sized channels allow the cells to be grown in a more biologically relevant environment, enabling cell to cell communications with continuous bathing in biological fluids in a tissue-like fashion. They also closely represent tissue and organ functionality by recapitulating mechanical forces as well as vascular perfusion. Here, we describe the use of humanized BBB model created with microfluidic organ-on-a-chip technology where human brain microvascular endothelial cells (BMECs) are cocultured with primary human pericytes and astrocytes. We thoroughly described the method to assess BBB integrity using a microfluidic chip and various sizes of labeled dextran as permeability markers. In addition, we provide a detailed protocol on how to microscopically investigate the tight junction proteins expression between hBMECs. Key words Blood-brain barrier, Permeability, In vitro models, Microfluidics, Organ-on-chip, Dextran

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Introduction A highly controlled microenvironment is required to promote the normal functioning of the central nervous system (CNS). The existence of a biological barrier at the blood to brain interface separating the brain from the rest of the body was established when Paul Ehrlich noticed that the brain tissue did not get stained by a peripherally infused dye [1]. This biological barrier is established by different cells at the interface between blood and brain tissue which is known as the blood-brain barrier (BBB). The BBB is

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formed by microvascular endothelial cells lining the cerebral capillaries penetrating the brain and spinal cord of most mammals and other organisms with a well-developed CNS [2]. The BBB plays a critical role in protecting the brain parenchyma from blood-borne agents and providing a major obstacle to the entry of drugs and other exogenous compounds into the CNS. It also forms a dynamic interface between blood and the CNS, thus controlling the influx and efflux of biological substances needed to sustain the brain metabolic processes and neuronal functions [3]. Unlike other vascular endothelial cells lining the peripheral blood vessels, brain microvascular endothelial cells present distinctive morphological, structural, and functional characteristics that set them apart from other vascular endothelia. These unique features include the expression of tight junction (TJ) proteins that seal the paracellular pathways between adjacent endothelial cells, thus preventing the unregulated passage of polar (water-soluble) molecules between the blood and the brain [4]. The TJs consist of three integral membrane proteins, namely, claudin, occludin, and junction adhesion molecules (JAMS), and several cytoplasmic accessory proteins including Zonula occludens-1, occludens-2, occludens-3 (ZO-1, ZO-2, ZO-3), cingulin, and others. The cytoplasmic proteins link membrane proteins to actin, which is the primary cytoskeleton protein for the maintenance of structural and functional integrity of the endothelium [2, 4]. In addition to microvascular endothelial cells, the capillary basement membrane (BM), astrocytes, and pericytes (PCs) embedded within the basal membrane, microglial and neuronal cells complete the structure of what is known as the neurovascular unit (NVU) [5]. The astrocytes and pericytes provide signals that are required for differentiation of the BMECs, and all three cell types are needed to maintain BBB integrity in vivo as well as in vitro [6]. For instance, in vitro studies have shown that ECs associated with PCs are more resistant to apoptosis than isolated endothelial cells, further supporting the role of PCs in supporting the structural integrity and genesis of the BBB [7]. The contribution of astrocyte cells in developing and maintaining the BBB has supported by several grafting and cell culture studies [8, 9]. An in vitro study showed that the optimal generation of BBB necessitates direct contact between endothelial cells and astrocytes [10]. The induction of BBB characteristics in ECs may be attributed not only to the NVU cells, but their cell-derived soluble factors as human or bovine endothelial cell monolayers showed a higher transendothelial resistance (TEER) when being cultured in astrocyte-conditioned media [11]. For studying the BBB, different animal models still provide the most reliable measurements for drug permeability considering the complex nature of the BBB function and structure; thus, they remain the gold standard for this purpose [12]. However, these models still have drawbacks such as high cost, low throughput, and

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labor-intensive [13]. The high demand for high-throughput methods that facilitate the screening of numerous compounds resulted in great interest in developing novel in vitro models. Recently, in vitro BBB models have been used widely owing to their improved reproducibility and low cost. In addition, they can be used for high-throughput screening and mechanistic studies [12]. Different cerebrovascular ECs derived from a variety of sources (primary, immortalized, and stem cell) can be used in a multitude of experimental setups [14–18]. To date, Transwells are considered the most valuable and widely used in vitro platforms for creating artificial BBBs. These models can be developed by culturing human, murine, or porcine brain-derived endothelial cells on a permeable membrane of an insert that is coated with an extracellular matrix [19]. The cells can be grown in a single monolayer or co-culture with astrocytes or other components of NVU [15]. The Transwell provides the structure that mimics the luminal and abluminal surfaces of the BBB. However, these models suffer from major drawbacks such as the lack of flow and shear stress, three-dimensional structure, and the presence of membrane between cells, which limits the ability to study the juxtracrine effect between cells. Shear stress is one of the most important but underestimated physiological stimuli, which contributes to vascular EC differentiation and maturation besides other cellular and molecular signaling. Akin to responses to inflammatory cytokines, shear stress has been shown to cause dramatic changes in EC morphology [20], gene expression [3, 21], and function [22]. That necessitates the development of an in vitro platform where we can apply shear stress. Organ-on-a-chip platforms offer 3D engineered microscale systems that can mimic the cellular microenvironment [23]. These systems closely represent tissue and organ functionality by recreating multicellular architectures, tissue-tissue interfaces, mechanical forces, physicochemical microenvironments, and vascular perfusion. The utility of organ-on-a-chip platforms was demonstrated by modeling different aspects of the BBB, including continuous luminal perfusion, real-time TEER monitoring, livecell imaging for permeability measurements, and metabolism pathways [3, 24–27]. Here, we utilized an enhanced human BBB model created with microfluidic organ-on-a-chip technology where we can culture human BMECs along with primary human pericytes and astrocytes (Fig. 1) [23, 28]. This platform also allows us to apply shear stress to these cocultured cells which have a critical role in BMEC maturation and mimic the vascular perfusion. We also used the platform to assess the BBB integrity by measuring the permeability of different molecular-sized labeled dextran and microscopically investigate the TJ proteins between the endothelial cells. Various BBB permeability markers are currently being used for in vitro studies, including sodium fluorescein and dextrans. Dextrans are complex, branched polysaccharides consisting of

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Fig. 1 Schematic diagram showing the different cells and compartments of the BBB chip; primary hBMECs seeded on the luminal side and primary human astrocytes and pericytes seeded on the abluminal side

many glucose molecules. They are commercially available labeled either with a fluorophore or biotin with their chain length varying from 3 to 2000 kDa [29]. In our protocol, we described the use of fluoro-labeled dextran as it is available in different molecular sizes, easy to visualize, and can be precisely quantified.

2

Materials Prepare all solutions using ultrapure water (double-distilled deionized water to attain a sensitivity of 18 MΩ cm at 25 °C) and analytical-grade reagents. Prepare and store all reagents at room temperature unless otherwise indicated. Please follow all waste disposal regulations when disposing of waste materials.

2.1

Reagents

1. Collagen IV. 2. Fibronectin. 3. Poly-L-lysine. 4. Dulbecco’s phosphate-buffered saline (DPBS). 5. 70% ethanol. 6. 4% paraformaldehyde. 7. Triton X-100. 8. Goat serum. 9. DMEM/F12. 10. Fetal bovine serum (FBS). 11. Fluorescein isothiocyanate (FITC)-dextran (3, 10, and 70 kDa). 12. Primary antibodies for Claudin-5, ZO-1, and Occludin.

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13. Alexa Fluor 488 and 555 secondary antibodies. 14. Human brain microvascular endothelial cells (hBMECs). 15. Human astrocytes (HA). 16. Human pericytes (HP). 17. Endothelial cell medium (ScienCell). 18. Astrocyte medium (ScienCell). 19. Pericyte medium (ScienCell). 2.2

Equipment

1. Two-layer microfluidic chip separated by a porous PDMS membrane. Chip dimensions: Top channel 1 mm × 1 mm and Bottom channel 1 mm × 0.2 mm (width × height). Porous PDMS membrane with 50 μm thickness and 7 μm pore size. 2. 15 mL conical sterile polypropylene centrifuge tubes. 3. 50 mL conical sterile polypropylene centrifuge tubes. 4. Serological pipettes. 5. Pipettes for volumes 10–1000 μL. 6. 150 mm cell culture dish. 7. T-75 flasks. 8. Centrifuge for cell culture. 9. BSC laminar flow hood. 10. CO2 gas incubator. 11. Cell counting chamber Neubauer hemocytometer. 12. Peristaltic pump. 13. 96-well clear-bottom black microplate. 14. Microplate reader. 15. Microscope.

3 3.1

Methods Cell Culture

1. Prepare media for each cell type as per vendor recommendation and keep it at 4 °C (see Note 1). 2. Prepare a fibronectin-coated culture vessel (3 μg/cm2, T-75 flask is recommended) for hBMECs. Prepare three to four separate flasks in order to have enough cells for multiple seeding. Coat the flasks and leave at 37 °C incubator overnight. 3. Prepare a Poly-d-Lysine-coated culture vessel (2 μg/cm2, T-75 flask is recommended) for astrocyte and pericyte subculture. Coat the flasks and leave at 37 °C incubator overnight.

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4. Thaw cells vial by briefly immersing in a 37 °C water bath. Carefully observe with gentle agitation and remove from the bath just before the last ice pellet melt. Avoid the complete melting of the vial content (see Note 2). 5. Spray the vial with 70% ethanol and dry it with before transferring to BSC laminar flow hood. 6. Transfer the content of the vial into 3 mL of prewarmed media of the respective cell in 15 mL sterile conical tube. Rinse the vial with an additional 1 mL to fully remove any residual cells and transfer to the 15 mL tube. 7. Bring the final volume to 15 mL with culture media and centrifuge at 500 rpm for 5 min at room temperature. 8. Carefully aspirate and discard supernatant, leaving a small media amount (100 μL) above the cell pellet to avoid cell aspiration. 9. Resuspend the cell pellet with 5 mL of warm culture media and gently pipet using 1 mL pipet to loosen the cell pellet (see Note 3). 10. Transfer the cell suspension to precoated T-75 flasks of the respective cell and add an extra 5–10 mL of warm media. Incubate overnight at 37 °C and 5% CO2. 11. The next morning, replace the culture media with a fresh prewarmed one. Continue changing the media every 2–3 days till the cells reach 70% confluency, and then replace every day until the culture is approximately 90% confluent. 3.2 Microfluidic Chip Preparation

1. Collagen/fibronectin coating solution: add 400 μL of collagen IV plus 100 μL of fibronectin solution (assuming that collagen IV and fibronectin stock concentration is 1 mg/mL) to 500 μL of sterile DPBS on a daily basis (see Note 4). Prepare 100 μL to each chip. 2. Using a 100 μL pipet, introduce the coating solution into the bottom channel inlet until a small droplet forms on the outlet. Without releasing the pipet plunger, take the pipet out from the bottom channel inlet and introduce the remaining solution into the inlet of the upper channel, leaving small droplets of excess coating solution on both inlets and outlets (see Note 5). 3. Place the chips inside a humid reservoir to provide extra humidity to avoid coating solution dryness and shrinkage. An empty 1 mL pipet tip container can be used. Add 5 mL of sterile DPBS to the bottom of the container and place the chips on the tip rack. 4. Close the container lid and incubate the chips at 4 °C overnight and then at 37 °C for 1 h the next day. For expedite coating, incubate the chips at 37 °C for 4 h.

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1. Trypsinize and harvest a culture flask containing the hBMECs. 2. A cell density of 1.5–2 × 107 must be adjusted before seeding the bottom channel. 3. Fully aspirate the coating solution from each channel. 4. Gently wash both channels with hBMEC culture media. Carefully aspirate the medium outflow leaving a small volume of excess medium covering the inlet and outlet ports. Channels should be kept filled with hBMEC culture medium (see Note 6). 5. Gently pipet hBMEC cell suspension to ensure a homogeneous cell suspension. 6. Carefully introduce 15–20 μL of hBMEC cell suspension with the optimized density into the bottom channel (see Note 7). 7. Check the chip under a microscope to ensure an even layer of cells is dispersed homogeneously along the length of the bottom channel (see Note 8). 8. Invert the chip to maximize cell attachment on the membrane (upper surface of the bottom channel). A 200 μL pipet tip can be used to level the inverted chips. 9. Place the seeded chip back into the humid reservoir and incubate at 37 °C for at least 4 h (see Note 9). 10. After 4 h, check the chip under a microscope to confirm that cells have attached to the upper surface (see Note 10). 11. Gently wash the bottom channel with 200 μL of warm hBMEC culture medium while aspirating the outflow as previously described. 12. Wash the top channel with prewarmed DMEM/F12 supplemented with 10% FBS culture medium while aspirating the outflow leaving the top channel filled with medium with small excess over the inlet and outlet ports. 13. Ensure there is no bubble in either channel, place them back in the reservoir, and incubate at 37 °C until cells are ready to seed again. 14. Trypsinize and harvest another culture flask containing the hBMECs and adjust the cell density as before. 15. Retrieve the washed chips from the incubator, and gently introduce 15–20 μL of hBMECs cell suspension into the bottom channel while aspirating the outflow as described before. 16. Without inversion, keep the chips into the reservoir back in the incubator until top channel cells are ready to seed. 17. Trypsinize and harvest astrocytes and pericytes from the culture flask and adjust the cell density to 2 × 106 and 0.2 × 106, respectively.

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18. Combine equal volumes of these cells to achieve a final coculture cell suspension density of 1 × 106 cells/mL of astrocytes and 0.1 × 106 cells/mL of pericytes (see Note 11). 19. Retrieve the chips from the incubator and carefully introduce 35–50 μL of the coculture cell suspension into the top channel while aspirating the outflow as previously described. 20. Check the chip under a microscope to ensure an even layer of cells is dispersed homogeneously along the length of the top channel. 21. Place the seeded chip back into the humid reservoir and incubate at 37 °C for at least 4 h or until cell attachment to the membrane. 22. After incubation, gently wash the top and bottom channels with 200 μL of respective media to remove unattached and dead cells (see Note 12). 23. Place the chips with inserted pipet tips into the reservoir, and incubate at 37 °C overnight at static conditions. 24. After overnight incubation, connect the seeded chips to a peristaltic pump and adjust the hBMEC medium flow rate in the bottom channel between 60 and 100 μL/h. Leave the chips under flow condition for 24 h to allow the BBB chips to adjust to the flow condition. 3.4 Measuring BBB Integrity 3.4.1

Permeability Assay

1. After adapting the chip to flow, barrier integrity can be assessed by calculating the apparent permeability of different molecular size dextran tracers. At this point, the chip is ready to start treatment and running the permeability assay after completion of the desired treatment period. 2. A working solution of 100 μg/mL of FITC-dextran tracers is prepared fresh in the hBMEC culture medium. 3. Prepare a standard curve for each tracer using serial concentrations in hBMEC culture medium. Place serial concentrations in 96-well plates and measure fluorescence intensity using the microplate reader. Set the excitation wavelength on the plate reader to 488 nm and the emission wavelength to 520 nm. Read the plate according to plate reader instructions. 4. Infuse the FITC-dextran solution through the bottom channel using the peristaltic pump at the optimized flow rate for 30, 60, and 120 min. Flow should be applied at the same rate for both top and bottom channels (Fig. 2). 5. Collect effluent samples from both channels and measure the fluorescent intensity of the samples (see Note 13). 6. Calculate the tracer concentration in each sample from the respective standard curve.

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Fig. 2 Schematic of BBB permeability assay, with tracer added to the luminal side media and samples collected with permeable tracer from the abluminal side

7. Use the following equation to calculate the apparent permeability (Papp): P app =

A×t ×

V r × Cr ðC d‐out × V d þC r × V r Þ ðV d × V r Þ

where Vr is the volume of the top channel (receiver) effluent at time t; Vd is the volume of dosing channel (bottom) at time t; A is the area of the membrane, which is 16.7 mm2 in this chip model; Cr is the measured tracer concentration in the top channel; and Cd-out is the measured tracer concentration in the dosing channel (bottom) effluent (see Note 14). 8. Blot the calculated Papp (cm/s) versus time of each tracer before and after treatment to assess barrier permeability. 3.4.2 Immunofluorescence Microscopy

1. Gently wash both channels with 200 μL of PBS thrice using 200 μL while aspirating the outflow at a slight distance from channel outlet to avoid cell detachment. 2. Fix the chips by adding 4% paraformaldehyde for 15 min (see Note 15). 3. Block with 10% goat serum in PBS with 0.2 Triton X-100 for 30 min (see Note 16). 4. Dilute primary antibody in 10% goat serum in PBS at 1:100 or as specified by the supplier. 5. Add 100 μL of primary antibody to the chip and incubate overnight at 4 °C. 6. On the next morning, wash thrice using 200 μL of PBS as previously described. 7. Add secondary antibody in 10% goat serum in PBS at 1:200 times dilution and incubate for 2 h at room temperature. Cover with aluminum foil and keep away from light.

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8. After 2 h, wash thrice with 200 μL of PBS as previously described. 9. Chips are ready to check under immunofluorescence or confocal microscope. 10. Transfer pictures to ImageJ to quantify fluorescence intensity at the cell-cell junctional interface.

4

Notes 1. Use within 15 days of preparation. You can aliquot to multiple 50 mL conical tubes to avoid multiple warm-cooling cycles. 2. Make sure the vial cap is tightly closed to avoid any liquid leakage inside the vial and keep the vial lid always above the water level. 3. Avoid many times pipetting as it may affect cell viability. 4. Maintain all components and mixture on the ice at all times and avoid bubble formation during pipetting. 5. Inspect each channel to ensure the bubble-free coating. If bubble is present, aspirate both channels and carefully introduce the solution again into each channel as previously described. 6. Place the aspirator nozzle slightly away from the outlet to avoid sudden aspiration of media inside the channel and bubble creation. 7. Aspirate the outflow at a slight distance from the outlet port to avoid cell aspiration from the channel. 8. If the seeding density is not optimal, wash the bottom channel twice with medium and reintroduce the cells after optimizing the cell density. 9. Avoid handling too many chips at once to minimize the amount of time cells remain outside the incubator. 10. Incubation time may need to be optimized to obtain maximum cell attachment. 11. Cell density can be optimized based on the proliferation rate and viability of cells. 12. Leave a filtered 200 μL pipet tip filled with respective media in each channel inlet and outlet ports to avoid medium mixing and keep enough medium for cells for overnight incubation. 13. Samples may need to be diluted to fall within the standard curve concentration limit. 14. Do not forget to consider the cumulative effect when calculating Papp for 60 and 120 min.

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15. Do not count the first time as the 4% PFA with being diluted with PBS remaining in the channel. 16. Store the blocking solution stock at 4 °C for 3 days and discard it after. References 1. Liebner S, Czupalla CJ, Wolburg H (2011) Current concepts of blood-brain barrier development. Int J Dev Biol 55(4–5):467–476 2. 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 3. Cucullo L, Hossain M, Puvenna V, Marchi N, Janigro D (2011) The role of shear stress in Blood-Brain Barrier endothelial physiology. BMC Neurosci 12:40 4. Ballabh P, Braun A, Nedergaard M (2004) The blood-brain barrier: an overview: structure, regulation, and clinical implications. Neurobiol Dis 16(1):1–13 5. Zlokovic BV (2011) Neurovascular pathways to neurodegeneration in Alzheimer’s disease and other disorders. Nat Rev Neurosci 12(12):723–738 6. Cecchelli R, Berezowski V, Lundquist S, Culot M, Renftel M, Dehouck MP et al (2007) Modelling of the blood-brain barrier in drug discovery and development. Nat Rev Drug Discov 6(8):650–661 7. Ramsauer M, Krause D, Dermietzel R (2002) Angiogenesis of the blood-brain barrier in vitro and the function of cerebral pericytes. FASEB J 16(10):1274–1276 8. Stewart PA, Wiley MJ (1981) Developing nervous tissue induces formation of blood-brain barrier characteristics in invading endothelial cells: a study using quail—chick transplantation chimeras. Dev Biol 84(1):183–192 9. Janzer RC, Raff MC (1987) Astrocytes induce blood-brain barrier properties in endothelial cells. Nature 325(6101):253–257 10. Rubin LL, Barbu K, Bard F, Cannon C, Hall DE, Horner H et al (1991) Differentiation of brain endothelial cells in cell culture. Ann N Y Acad Sci 633:420–425 11. Neuhaus J, Risau W, Wolburg H (1991) Induction of blood-brain barrier characteristics in bovine brain endothelial cells by rat astroglial cells in transfilter coculture. Ann N Y Acad Sci 633:578–580 12. 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 13. van der Helm MW, van der Meer AD, Eijkel JC, van den Berg A, Segerink LI (2016) Microfluidic organ-on-chip technology for bloodbrain barrier research. Tissue Barriers 4(1): e1142493 14. Faria A, Pestana D, Teixeira D, Azevedo J, De Freitas V, Mateus N et al (2010) Flavonoid transport across RBE4 cells: a blood-brain barrier model. Cell Mol Biol Lett 15(2):234–241 15. Hatherell K, Couraud PO, Romero IA, Weksler B, Pilkington GJ (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 16. 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 Mol Biol 814:415–430 17. Paolinelli R, Corada M, Ferrarini L, Devraj K, Artus C, Czupalla CJ 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 18. 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 19. Helms HC, Abbott NJ, Burek M, Cecchelli R, Couraud PO, Deli MA 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 20. Ott MJ, Olson JL, Ballermann BJ (1995) Chronic in vitro flow promotes ultrastructural differentiation of endothelial cells. Endothelium 3(1):21–30 21. Akimoto S, Mitsumata M, Sasaguri T, Yoshida Y (2000) Laminar shear stress inhibits vascular endothelial cell proliferation by inducing cyclin-dependent kinase inhibitor p21(Sdi1/ Cip1/Waf1). Circ Res 86(2):185–190

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22. Ngai AC, Winn HR (1995) Modulation of cerebral arteriolar diameter by intraluminal flow and pressure. Circ Res 77(4):832–840 23. Bhatia SN, Ingber DE (2014) Microfluidic organs-on-chips. Nat Biotechnol 32(8): 760–772 24. Booth R, Kim H (2014) Permeability analysis of neuroactive drugs through a dynamic microfluidic in vitro blood-brain barrier model. Ann Biomed Eng 42(12):2379–2391 25. Cho H, Seo JH, Wong KH, Terasaki Y, Park J, Bong K et al (2015) Three-dimensional bloodbrain barrier model for in vitro studies of neurovascular pathology. Sci Rep 5:15222 26. Maoz BM, Herland A, FitzGerald EA, Grevesse T, Vidoudez C, Pacheco AR et al (2018) A linked organ-on-chip model of the human neurovascular unit reveals the

metabolic coupling of endothelial and neuronal cells. Nat Biotechnol 36(9):865–874 27. Prabhakarpandian B, Shen MC, Nichols JB, Garson CJ, Mills IR, Matar MM et al (2015) Synthetic tumor networks for screening drug delivery systems. J Control Release 201:49–55 28. Park TE, Mustafaoglu N, Herland A, Hasselkus R, Mannix R, FitzGerald EA et al (2019) Hypoxia-enhanced blood-brain barrier chip recapitulates human barrier function and shuttling of drugs and antibodies. Nat Commun 10(1):2621 29. 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

Chapter 8 Intravital Imaging of Leukocyte-Endothelial Interaction in Hindlimb Ischemia/Reperfusion Injury by Intravital Multiphoton Microscopy Johannes Zeller, Karlheinz Peter, and Steffen U. Eisenhardt Abstract Ischemia/reperfusion injury in skeletal muscle leads to sterile inflammation and affects structure and function permanently. However, the main understanding of the molecular and cellular mechanisms mainly relies on in vitro and ex vivo investigations. Recent advances in intravital microscopy allow for insights into dynamic processes at the cellular and subcellular level under both physiological and pathophysiological conditions. Real-time intravital imaging by two-photon microscopy (2P-IVM) has emerged as a powerful tool in the evaluation of the cell-cell interaction and molecular biology of leukocytes in live animals. Acute ischemic injury in limbs may occur due to crush syndrome, compartment syndrome, and vascular diseases and injury as in acute peripheral arterial occlusion, caused by a diverse array of pathological conditions. Iatrogenic revascularization and restoration of perfusion results paradoxically in aggravated tissue injury. Furthermore, the effects of IR-injured skeletal muscle in clinical conditions such as compartment syndrome or crush syndrome may induce rhabdomyolysis and are associated with so-called remote injuries as acute kidney dysfunction. Here, we discuss the considerations for and describe a 2P-IVM method designed for visualization of leukocyte-endothelial interaction. This chapter will provide a detailed experimental setup and a step-by-step protocol for the dynamic imaging of leukocyte-endothelial-interaction in an ischemia/ reperfusion injury model. Key words Two-photon intravital microscopy, Ischemia/reperfusion injury, Leukocyte-endothelial interaction

1

Introduction

1.1 Why Use Multiphoton Excitation in Intravital Microscopy?

Intravital imaging by multiphoton microscopy is a powerful tool in the evaluation of the molecular biology in leukocyte interaction with vascular endothelium. Two-photon excited fluorescence (TPE) provides for major advantages over single-photon excitation fluorescence intravital microscopy with epifluorescence wide-field microscopes and confocal microscopes: (1) Due to near-infrared excitation wavelength in TPE, tissue damage by photo-toxicity is reduced, with energy per photon Ephoton = h × f and thus,

Binu Tharakan (ed.), Vascular Hyperpermeability: Methods and Protocols, Methods in Molecular Biology, vol. 2711, https://doi.org/10.1007/978-1-0716-3429-5_8, © Springer Science+Business Media, LLC, part of Springer Nature 2024

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equivalently, inversely proportional to the wavelength. In other words, the longer the photon’s wavelength, the lower its energy. (2) Second, longer wavelength results in less scattering and therefore greater tissue penetration with up to a range of a few millimeters. The strong wavelength dependence of the scattering (~λ4) is, interestingly, something we observe everyday: shorter (blue) wavelengths are scattered more strongly than longer wavelengths (yellow and red), giving the sky its blue color at daytime and the reddish hue of the low sun. (3) The focal plane is created by “confocality” of the laser beam, not by a detector pinhole. Hence, the signal-to-noise ratio (SNR) is improved and phototoxicity is reduced. These properties make TPE a most useful tool for intravital imaging of molecular biological processes. 1.2 Considerations for the Microscope Setup

In general, there are two builds of microscopes used for multiphoton microscopy: inverted microscopes and upright microscopes. While inverted microscopes are advantageous for in vitro live cell imaging, upright builds are more convenient in intravital microscopy, especially with dipping objective lenses. The preferable choice would be an upright microscope when working with animals for intravital imaging. First, the microscope must be established in a quiet and dark room. An optical table will provide a stable and shock-proof basis for your microscope. Second, the surrounding can have a major influence on the accuracy of your imaging. Therefore, it is important that the interior of your imaging facility can be completely darkened. If this is not feasible, consider a custom-made tent for your microscope, e.g., made of item building kit and lightproof fabric. Room temperature should remain relatively stable, and the microscope should obtain optimal operating temperature before each experiment. Third, provide sufficient personal protection. Consider warning lights, when the laser is running, laser protection goggles for the experimenter, and safety instruction by a laser safety designate.

1.3 Why Use the Tibialis Anterior Muscle for the Evaluation of Leukocyte-Endothelial Interaction?

The capillary bed in the muscle tissue represents a convenient imaging field for direct visualization and observation of leukocyte cell-cell interaction under normal and pathologic conditions. Healthy skeletal muscle provides a dense capillary bed and hosts only exceedingly rare resident leukocytes; thus, alterations of leukocyte count in the interstitial space are unambiguous and can serve as an appropriate surrogate for inflammation [1, 2]. The cremaster muscle and tibialis anterior muscle are two of the most commonly used muscle tissues in intravital imaging [3]. The cremaster muscle covers the testes, and exteriorization has to be performed before imaging is possible [3]. Therefore, preparation might be difficult and prone to experimenter’s experience and skills. Surgical manipulation to the muscle itself may induce significant inflammation, thus distorting the baseline of resting muscle tissue. The tibialis

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anterior muscle of the hindlimb presents a more conveniently observable microcirculation with easy access, therefore low variability and high reproducibility in the visualization of leukocyteendothelial interaction. However, the tibialis anterior muscle model described hereafter is more prone to interfering artifacts from the breathing motion. Hence, we present a feasible method to restrain the hindlimb without hampering the microcirculation. The extent of tissue damage inflicted by ischemia and reperfusioninjury depends on various factors, with critical ischemia time, temperature, and tissue type of particular importance. Muscle susceptibility to IRI depends on the muscle fiber type. Muscles are composed of either mainly glycolytic or oxidative fibers [4]. In the rat hindlimb, the superficial portion of the tibialis anterior muscle represents a type IIb muscle, a muscle composed mainly of fast-twitch glycolytic fibers for locomotion [5]; thus, it is less resilient to ischemia and more prone to the ischemia/reperfusion injury.

2

Materials

2.1 Preparation of the Tracer: Labeling Bovine Serum Albumin with Texas Red®

Bovine serum albumin (BSA) conjugated with fluorescent dyes like Texas Red® is commonly used as a tracer for clear delineation of the microcirculation in intravital microscopy. The molecular weight of BSA is approximately 66,000 Da, which allows for differentiation of the blood vessel area from interstitial space area even under inflammatory conditions. Labeling of BSA with Texas Red®-maleimide represents a cost-effective way. Maleimide shows high selective affinity toward thiols (cysteine residues in polypeptides and proteins). Aim for a fluorophore-to-protein value (molar ratio) of 10 moles Texas Red® per 1 mole BSA. 1. Prepare 1 M sodium bicarbonate solution. 2. Dissolve BSA in degassed Dulbecco’s PBS 500 mg/100 mL at pH 7.4. 3. Dissolve Texas Red® maleimide in DMSO 1–10 mg per 100 μL. 4. Add DTT to reduce the disulfide bonds. Keep at room temperature. 5. Check the pH of the BSA solution and adjust it if necessary to pH 8.5–9.5 by adding 1 M sodium bicarbonate. 6. The total volume of the Texas Red® maleimide solution in DMSO has to be less than 10% of the total reaction volume. 7. Test your Texas Red®-conjugated BSA and adjust if necessary (see Note 1).

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Animals

1. Transgenic CD68-GFP Wistar rats, 180–220 g body weight. 2. Alternatively, male Wistar rats, 180–220 g body weight.

2.3

Supplies

1. Isoflurane, e.g., Abbott, Wiesbaden, Germany. 2. Vaporizer for isoflurane. 3. Plexiglas box to accommodate rats. 4. Buprenorphine, Buprenex or equal. 5. Eye lubricant, e.g., Bepanthen® eye and nose cream. 6. Heating table or lamp, a thermometer for rectal temperature measurement. 7. Non-alcoholic disinfectant, e.g., Braunol® 7.5% povidoneiodine, B. Braun, Melsungen, Germany. 8. Syringes, 1 and 5 mL. 9. 27G cannula for subcutaneous application of pain killer. 10. Abbocath-T intravenous cannula 26G, 0.6 × 19 mm. 11. Sterile normal saline (0.9% NaCl solution). 12. Gauze sponges. 13. Medical tape, e.g., Leukopor® micropore paper tape. 14. Prewarmed ultrapure deionized water. 15. Water bath at 37 °C. 16. Incubator at 37 °C. 17. Heat/cooling packs. 18. Styrofoam box for insulation. 19. Lens cleaning utensils.

2.4 Optional: For Heart Rate Monitoring

1. Abbocath-T intravenous cannula 26G, 0.6 × 19 mm. 2. Three-way valve, e.g., Discofix® C, B. Braun, Melsungen, Germany. 3. Tubing Type IV-standard, Luer. 4. Pressure monitoring system, e.g., smith medical Logical® MX960 and Logical® (MX960XY), MEDEX™ medical Ltd.

2.5 Surgical Instruments

All surgical instruments have to be clean and sterilized (by autoclave or chemiclave) before every use. 1. Fine dissecting scissors, curved Jameson or Metzenbaum scissors or equal. 2. Fine Adson forceps, surgical (with teeth). 3. Bovie® electric cautery. 4. Micro-scissors, straight and curved Adventitia scissors. 5. Micro-forceps.

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6. Micro-vessel clamp, Biemer or equal. 7. Micro-vessel clamp applicator. 8. Bulldog clamp. 2.6 Microscope Setup for Two-Photon Intravital Microscopy of I/R Injured Muscle

3

Two-photon intravital imaging of anaesthetized rats was performed using an upright Zeiss LSM 980 microscope equipped with Airyscan 2, LED light source for fluorescence microscopy Colibri 7 system (UV-line 385/30 nm, B-line 469/38 nm, and G-line 555/30 nm), and a Chameleon Discovery wide-range tunable laser system (Coherent Inc., Santa Clara, CA, USA) (see Note 2).

Methods

3.1 Tail Vein Cannulation for Intravenous Injections

For the intravenous application of fluorescent dyes, saline, and drugs of interest, perform a venous cannulation. The tail vein cannulation represents an easy access to the circulation with only minor invasivity and a steep learning curve (see Note 5). 1. Anaesthetize the rat. Sit the rat in the Plexiglas box and introduce 2.5% v/v isoflurane for introduction of anesthesia. For maintenance reduce isoflurane to 1.5% (see Note 6). 2. Place the rat in a prone position on the heating pad with its tail dangling over the edge of the table. Hydrostatic pressure will fill the tail veins passively and thus facilitate the cannulation (see Notes 7 and 8). 3. If tail veins are still only slightly visible, it is helpful to place the tail into tepid water for approx. ½ minute or warm it with a warming lamp. 4. Hold the tail between your thumb and middle finger and lay it over your extended index finger, so that the tail is laying straight between thumb and index finger (Fig. 1a). 5. Rinse an intravenous catheter (Abbocath-T 26 G, 0.6 × 19 mm) with sterile saline. The Luer ending of your cannula should be clear and filled with saline solution. A small trace of blood within the saline can clearly indicate the correct positioning of your cannula (Fig. 1b). 6. Puncture the vein through the skin in a flat angle (Fig. 1a). The skin is sturdy; thus, repeated use of one needle is not recommended. 7. Remove the needle (Fig. 1c). 8. Test your access for patency with sterile saline and fix it with medical tape. 9. Flush a three-way valve with prewarmed sterile saline solution, and connect it to the fixed intravenous catheter.

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Fig. 1 Tail vein cannulation. (a) The rat is a lateral position; hold the firmly between your index finger and thumb. The cannula enters the skin in a 45-degree angle. (b) Blood enters the clear ending of the intravenous catheter as a sign of correct positioning. (c, d) Remove the cannula and fix the plastic tube with medical tape. Connect the three-way valve. Test for correct intravenous position: if saline solution can be injected without force and swelling of the tail, the catheter position is correct

10. Add a 5 mL syringe filled with saline on one end of the valve (Fig. 1d). 11. The free opening is reserved for the application of drugs or fluorescent dyes. 3.2 Tail Artery Cannulation for Monitoring of Blood Parameters (Optional)

Measurement of blood parameters serves as a reliable monitoring for stress during surgery and imaging. We use the technique described by Hagmu¨ller et al. [6] with minor modifications by our workgroup. The following preparation steps are best accomplished using a surgical microscope and should be performed before preparing the animal for imaging. The cannulation of the tail artery provides a less invasive procedure compared to the cannulation of the carotid artery (see Note 9). 1. Prepare your tubing system at 37 °C. Warm all tubing, the three-way valve, and solutions to 37 °C, and thoroughly fill the tubing system with prewarmed saline solution. Check for small

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bubbles. The whole system has to be absolutely gas-free to correctly transduce the arterial pressure. Keep all components warmed in a Styrofoam box with warmed heat packs. 2. Under anesthesia, make a small longitudinal incision using a scalpel no. 15 slightly paramedian and on the ventral aspect of the tail approximately 1–2 cm distal from the tail’s base. 3. For the cannulation of the arteria caudalis mediana, separate the vessel by blunt dissection from the surrounding tissue. The artery runs in a groove in the midline of the tail under taut connective tissue. 4. Pass two pieces of equally long suture beneath the artery. The distal suture is for ligation of the artery. The proximal suture is yet only tied loosely around the artery. Use a micro-vessel clamp proximal to the loose suture. 5. Make a small incision into the tail artery between the two sutures near the distal ligature. 6. Insert the polyimide catheter (0.28 mm inner diameter) filled with saline and heparin; the liquid column will serve as a pressure transducer. 7. The polyimide tubing is connected to a three-way valve over Luer to a LogiCal® pressure monitoring system or equal. 8. Push the catheter forward to the base of the tail. After passing the loosely knot suture, tighten the knot. Now the clamp can be removed without bleeding. 9. Tie a second loop around the catheter and the artery to secure the catheter. Additional anchorage will prevent kinking. 10. Secure all tubing with medical tape. 11. Arterial blood gases can be monitored during the procedure by sampling small blood probes via the three-way valve and by using a blood gas analyzer [7]. Blood pH can be taken in consideration and may exceed physiological limits of 7.35–7.45 during the reperfusion phase of the experiment. 12. The heart rate can be monitored and used as an exclusion criteria for rats that show heart rate 360 bpm, as well as a mean arterial pressure dropping below 80 mmHg for longer than 5 min [8, 9]. 3.3 Surgical Procedure

1. Prepare a Plexiglas box with paper towels. Determine the tare weight of the box. 2. Gently place your test animal inside the box and weigh its body using an electronic scale. 3. Anaesthetize the test animal with 2.5% v/v volatile isoflurane for introduction and 1.5% v/v for maintenance. 4. Use eye ointment to prevent postoperative blinding.

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5. Place the rat onto the heating pad and maintain hypnosis via a silicon mask or cone. 6. Place a subcutaneous depot of 0.05 mg/kg bodyweight buprenorphine. The total volume should be adequate to the size of the test animal. For instance, rats weighing 180–220 g receive 300 μL volumes [10] for pain relief. 7. Use the toe pinch test and breathing pattern and rate to assess depth of anesthesia. If no reaction to the pinch occurs, proceed with the following. 8. Place the rat in prone position onto a temperature-controlled heating pad. 9. Shave one hindlimb carefully with a clipper. Clean the hindlimb thoroughly with wet paper towels or rinse the hindlimb under running water. Ensure you removed all loose hair. Otherwise, it will affect the quality of the imaging (see Note 3). 10. Use non-alcoholic disinfectant on the intact hindlimb, e.g., Braunol®. 11. All following steps are best accomplished using a surgical microscope. 12. Place the rat in a supine position and make a small oblique skin incision with the Jameson scissors 1 cm distal to the inguinal ligament (IL) (Fig. 2a, dotted black line). This structure demarcates the proximal border of the hindlimb to the abdominal wall and is well palpable through the intact skin. The femoral artery enters the thigh from behind the IL (Fig. 2a, dotted red line). Here, it is covered by the inguinal fat tissue (Fig. 2b). Detach the fat pad from the abdominal wall by blunt dissection. 13. Place the fat tissue flap to the distal part of the leg without disturbing the pedicle. Deep to the fat pad, the femoral artery is apparent (Fig. 2c). 14. Carefully dissect the artery from the surrounding tissue, the femoral nerve, and vein in blunt technique. First, make a small incision longitudinal to the vessels in the surrounding connective tissue (Fig. 2d). 15. Use a micro-vessel clamp and place it onto the femoral artery (Fig. 2e). Control sufficient occlusion by performing an emptyand-refill test distal to the clamp. If the artery stays collapsed after stroke out, occlusion is sufficient. Start your timer for the 90-min ischemia period. 16. Close the skin wound temporary with a bulldog clamp. And cover the rat with clean fabric to further maintain body temperature.

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Fig. 2 Preparation of the femoral artery for temporal occlusion. Deeper to the IFP, (a) the anaesthetized rat is in supine position; the hindlimb chosen for I/R injury is shaved and abducted. The border between hindlimb and abdominal wall (dotted black line) is visible and represents the inguinal ligament. The femoral artery and vein (dotted red and blue line, respectively) enter the thigh from behind the inguinal ligament. In the upper part of its course, the artery is medially related to the femoral vein and laterally to the femoral nerve and its branches. (b) Make a small skin incision along the black dotted line. Directly under the skin, the inguinal fat tissue covers the femoral vessels. (c) After detaching the inguinal fat pad from its attachment point to the lateral abdominal wall in blunt dissection, the inguinal ligament (white structure) and the femoral vessels are visible. (d, e) The vessels are accompanied by the femoral nerve. The bigger, more caudal structure is the femoral vein. Dissect the artery from the nerve and vein gently and clamp the artery with a micro-vessel clamp

17. After 80 min, turn the rat in a lateral position with the hindlimb of interest facing upward (Fig. 3a). 18. Make a small incision anterior to the lateral saphenous vein by lifting the skin carefully with surgical forceps. The fascia connecting the gastrocnemius and the tibialis anterior muscle is visible through the skin (Fig. 3a, b, dotted line). Cut the skin longitudinal to the leg ~2.5 cm length. Bluntly separate the skin from the muscle fascia beneath (Fig. 3c). 19. Bleeding during ischemia should be minimal, and if it occurs use a saline-soaked gauze to perform subtle hemostasis.

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Fig. 3 Preparation of the tibialis anterior muscle for imaging. Depicted are the surgical steps for visualization of the tibialis anterior muscle. (a) The anaesthetized rat is in a lateral position. A groove between the gastrocnemius muscle and tibialis anterior muscle is visible through the skin. (b) The contour of the connecting fascia of both muscles is marked (dotted black line). (c) Make a skin incision along the dotted line. The tibialis anterior muscle is now visible ventral to the fascia. Both the fascia and the skin show a dense vessel bed. (d) Position the rat in prone position, abduct the hindlimb, and fix it within the leg restrainer. (e) Place the rat laying on the heating pad under the microscope. Evaluate body temperature during the imaging procedure using an anal thermometer

Bleeding from the skin can be stopped using a high temperature cautery (e.g., Bovie® electric cautery) sparingly. 20. Keep the fascia and the muscle tissue moist with prewarmed saline solution as soon as the skin is removed and muscle fascia is exposed. 21. Place the custom-made leg restrainer on the heating pad. Wait until it adopts the temperature from the temperaturecontrolled heating pad. 22. After 90 min, remove the micro-vessel clamp and wait for sufficient patency of the femoral artery. 23. Place the hindlimb in the hindlimb restrainer. Use a plastically deformable adhesive (e.g., Bostik Blu-Tack) if a size mismatch occurs, and model it semi-circumferential around the leg so

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Fig. 4 Setup for a in vivo imaging facility. The upright microscope for intravital imaging sits on an optical table. This provides for a stable and shock-proof basis and artifact-free imaging. The interior of the imaging facility can be completely darkened (see Note 4). If this is not feasible, consider a custom-made tent for your microscope, e.g., made of item building kit and lightproof fabric. Warning signs are added for additional safety

that the exposed tissue is placed under the window of the restrainer (Fig. 3d). 24. Adjust the lid with slight pressure onto the hindlimb without disturbing the circulation. 25. Fill the reservoir above the glass with prewarmed ultrapure deionized water. 26. Place the animal under the microscope and perform imaging procedure (Figs. 3e and 4). 3.4 Choosing Good Regions of Interest and the Right Vessel Diameter

Central for good reproducibility is to compare similar vessels: arteries, capillaries, and venules differ from each other in anatomy and diameter. 1. Locate an area with parallel capillaries within your muscle by screening it with epifluorescence excitation and the eyepiece. Turn the power of your LED system as low as possible to prevent photo-irritation and fading of your dyes. Adjust your eyes to the dark before using the eyepiece of the microscope. 2. To visualize the bloodstream in vivo, use Texas Red®-BSA or similar as a tracer.

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3. Immediately after, the tracer injection arteries and arterioles are delineated from surrounding tissue, followed by capillaries and post-capillary venules. This allows for differentiation in the field of view. 4. For repeated measurements in different test animals, the diameter of the post-capillary venules can be calculated with the fullwidth half-maximum (FWHM) method as described by [11]. The method assists in acquiring accurate measurements and comparability. 3.5 Digital Image and Video Analysis

1. Use image/video analysis software that allows for drawing in scale bars and play video sequences in slow motion, e.g., Zeiss ZEN 2 blue Imaging Software. 2. For the quantification of leukocyte interaction with the vascular endothelium, define a consistent length of a vessel section. Best, search for post-capillary venules, and determine the interval for all experiments, e.g., 20 s (see Notes 10 and 11).

4

Notes 1. Labeling of BSA can result in either strong or weak labeling. Check the degree of labeling using a DOL calculator for the absorbance of BSA (280 nm) and Texas Red® (586 nm). (“Quest Calculate™ Degree of Labeling (DOL) Calculator.” AAT Bioquest, Inc, 17 Jun. 2020, https://www.aatbio.com/ tools/degree-of-labeling-calculator). 2. Two-photon excitation differs in wavelength from familiar numbers for frequently used fluorophores and dyes. The Spectra Database for dyes hosted at the University of Arizona (http://www.spectra.arizona.edu/) provides information for considerate decision-making. 3. Rodent hair strongly absorbs and scatters light in both the visible and the near-infrared spectrum. Thus, thorough shaving of the test animal and removal of all loose hair are recommended to minimize autofluorescence and hampering of the imaging signals. 4. Darken the room beforehand and let your eyes adapt. When in “locate” mode, use as little light as possible to avoid photobleaching of your dyes and phototoxicity. 5. Check that your animal studies you perform are in compliance with the local ethical regulations. If possible, consult your local Animal Welfare Officer. All procedures should follow approved institutional and governmental animal procedures.

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6. Ensure proper anesthesia. Breath rate and breathing pattern are feasible surrogate parameters and next to the toe pinch test. Normopnoea of the rat is around 85 breaths per minute. 7. Avoid hypothermia or hyperthermia (35.9–37.5 °C) of the test animal. Rodents are not able to maintain body temperature under anesthesia. Sufficient body temperature control is therefore of utmost importance. 8. Sufficient fixation of your test animal is essential. In case of an upright microscope, use a window design as presented in Fig. 5. Feel free to download the blueprint for the chamber (https://www.uniklinik-freiburg.de/plastischechirurgie/ forschung/intravital-imaging-facility.html) or contact the author for information (Figs. 6 and 7). 9. Ensure that the blood sampling does not exceed 10% of the total blood volume (total blood volume is approximately 64 mL/kg body weight × 0.4 kg) [12]. The National Centre for the Replacement, Refinement & Reduction of Animals in Research provides further information and more defined

Fig. 5 Custom rat hindlimb restraint. Breathing motion is one of the main sources of artifacts in high-resolution intravital imaging. To minimize respiration-induced distortion, we developed a noninvasive external fixation for the hindlimb of small rodents. Please contact the authors for a blueprint or find it on https://www.uniklinikfreiburg.de/plastischechirurgie/forschung/intravital-imaging-facility.html

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Fig. 6 Leukocyte-endothelial interaction in tibialis anterior muscle. Transgenic CD68-GFP rats were used. Texas Red®-BSA serves as a reliable tracer for imaging microcirculation in inflammatory settings. GFP-positive CD68 cells can be detected over time (a) transmigrating into the muscle or in interaction with the vessel endothelium (b)

instructions on this issue: https://www.nc3rs.org.uk/ratdecision-tree-blood-sampling. 10. For profound statistical analysis of the acquired data, we suggest using spreadsheet software, e.g., Microsoft Excel, and a scientific graphing and statistic software GraphPad Prism 8. Obtain statistical advice by your institution’s statistic department before performing the animal experiments. It is recommended to perform statistical power analyses before planning animal experiments to comply with the principle of reduction in animals in research. Those statistics can be computed using G*Power software [13]. 11. Vascular permeability under basal conditions and vascular hyperpermeability under inflammatory conditions vary, and thus, molecular weight of the tracer (dextran, BSA or other) should be taken in consideration [14].

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Fig. 7 Muscle anatomy of the rat hindlimb. Shown is the rat hindlimb as a crosssection scheme. Tibialis anterior (TA) muscle can be differentiated into a glycolytic portion (type IIb, TAw) and deeper portion of fast twitching oxidativeglycolytic fibers (type IIa, TAr). The fast twitch glycolytic fibers are more prone to ischemia-reperfusion induced injury. Muscle fiber type composition according to Armstrong and Phelps [5]

Acknowledgments This work was supported by a grant to Steffen Eisenhardt by the German Research Foundation (DFG) (INST 39/1137-1FUGG) for a Multiphoton microscope. Steffen Eisenhardt is a Heisenberg Professor of the German Research Foundation (EI 866/4-1). We are grateful for expert advice and assembly of the hindlimb restraint from Gerd Strohmeier and Waldemar Schimpf, Central scientific workshops of the Neurocenter, Medical Center – University of Freiburg, Freiburg, Germany. We thank the expert advice of Animal Welfare Officer of the University Medical Center Freiburg Dr. Caroline Johner for her consultation. References 1. Hoppeler H, Mathieu O, Weibel ER et al (1981) Design of the mammalian respiratory system. VIII. Capillaries in skeletal muscles. Respir Physiol 44:129–150. https://doi.org/ 10.1016/0034-5687(81)90080-3 2. Welling TH, Davidson BL, Zelenock JA et al (1996) Systemic delivery of the interleukin-1

receptor antagonist protein using a new strategy of direct adenoviral-mediated gene transfer to skeletal muscle capillary endothelium in the isolated rat hindlimb. Hum Gene Ther 7: 1795–1802. https://doi.org/10.1089/hum. 1996.7.15-1795

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3. Thiele JR, Goerendt K, Stark GB, Eisenhardt SU (2012) Real-time digital imaging of leukocyte-endothelial interaction in ischemiareperfusion injury (IRI) of the rat cremaster muscle. J Vis Exp:e3973. https://doi.org/10. 3791/3973 4. Gillani S, Cao J, Suzuki T, Hak DJ (2012) The effect of ischemia reperfusion injury on skeletal muscle. Injury 43:670–675. https://doi.org/ 10.1016/j.injury.2011.03.008 5. Armstrong RB, Phelps RO (1984) Muscle fiber type composition of the rat hindlimb. Am J Anat 171:259–272. https://doi.org/10. 1002/aja.1001710303 6. Hagmu¨ller K, Liebmann P, Porta S, Rinner I (1992) A tail-artery cannulation method for the study of blood parameters in freely moving rats. J Pharmacol Toxicol Methods 28:79–83. https://doi.org/10.1016/1056-8719(92) 90051-2 7. von Dobschuetz E, Pahernik S, Hoffmann T et al (2004) Dynamic intravital fluorescence microscopy—a novel method for the assessment of microvascular permeability in acute pancreatitis. Microvasc Res 67:55–63. https://doi.org/10.1016/j.mvr.2003.09.006 8. Takasu A, Minagawa Y, Ando S et al (2010) Improved survival time with combined early blood transfusion and fluid administration in uncontrolled hemorrhagic shock in rats. J Trauma Inj Infect Crit Care 68:312–316.

h t t p s : // d o i . o r g / 1 0 . 1 0 9 7 / T A . 0b013e3181c48970 9. Vutskits L, Briner A, Klauser P et al (2008) Adverse effects of methylene blue on the central nervous system. Anesthesiology 108:684– 6 92 . h t t p s : //d o i . o r g / 1 0 . 1 0 9 7/ A L N . 0b013e3181684be4 10. Curtin LI, Grakowsky JA, Suarez M et al (2009) Evaluation of buprenorphine in a postoperative pain model in rats. Comp Med 59: 60–71 11. Driscoll JD, Shih AY, Drew PJ et al (2013) Two-Photon Imaging of Blood Flow in the Rat Cortex. Cold Spring Harb Protoc 2013: pdb.prot076513. https://doi.org/10.1101/ pdb.prot076513 12. Diehl K-H, Hull R, Morton D et al (2001) A good practice guide to the administration of substances and removal of blood, including routes and volumes. J Appl Toxicol 21:15–23. https://doi.org/10.1002/jat.727 13. Faul F, Erdfelder E, Lang A-G, Buchner A (2007) G*Power 3: a flexible statistical power analysis program for the social, behavioral, and biomedical sciences. Behav Res Methods 39: 1 7 5 – 1 9 1 . h t t p s : // d o i . o r g / 1 0 . 3 7 5 8 / BF03193146 14. Egawa G, Nakamizo S, Natsuaki Y et al (2013) Intravital analysis of vascular permeability in mice using two-photon microscopy. Sci Rep 3:1932. https://doi.org/10.1038/srep01932

Chapter 9 Studying Angiogenesis Using Matrigel In Vitro and In Vivo Anantha K. Kanugula, Ravi K. Adapala, Brianna D. Guarino, Neha Bhavnani, Harshitha Dudipala, Sailaja Paruchuri, and Charles K. Thodeti Abstract Angiogenesis plays a critical role in physiology and pathophysiology of the human body; hence, it is important to explore the methods to study angiogenesis under in vitro and in vivo settings. Here, we describe three different methods to assess angiogenesis using Matrigel: an in vitro two- or threedimensional (2D/3D) tube formation or angiogenesis assay using endothelial cells with growth factor supplemented Matrigel, an ex vivo sprouting angiogenesis assay embedding aortic rings in the Matrigel, and finally, Matrigel plug assays wherein Matrigels are implanted into the flanks of mice to assess the recruitment of endothelial cells to form new blood vessels in vivo. Key words Two-dimensional angiogenesis, Matrigel, Endothelial cells, Aortic rings

1

Introduction Angiogenesis, formation of new blood vessels from pre-existing ones, is a tightly regulated process involving various cell types and biomechanical signaling pathways. During physiological processes, angiogenesis is highly organized; however, it is disturbed under pathological conditions, for instance, cancer, diabetes, ischemic heart disease, arthritis, and psoriasis [1, 2]. Though angiogenesis refers to vascular proliferation, indicating a definite process, the reality is that angiogenesis is different in each pathological condition where uniform blood vessels are not generated [1, 2]. Thus, understanding the key factors and processes involved in the vessel formation and maturation is essential for the therapy of angiogenesis-dependent diseases. Angiogenic processes are coordinated by proangiogenic factors such as vascular endothelial growth factor (VEGF), basic fibroblast growth factor (bFGF), and antiangiogenic factors such as thrombospondins (TSP), angiostatin, and endostatin [3, 4]. These factors together with downstream signaling pathways govern the sprouting of neovessels. Therefore, it is

Binu Tharakan (ed.), Vascular Hyperpermeability: Methods and Protocols, Methods in Molecular Biology, vol. 2711, https://doi.org/10.1007/978-1-0716-3429-5_9, © Springer Science+Business Media, LLC, part of Springer Nature 2024

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important to study angiogenesis in depth and, more specifically, in the appropriate tissue and disease state to identify novel therapeutic strategies. One of the most important aspects in studying angiogenesis is to choose the most suitable assay. Besides the available variety of in vitro and in vivo assays, a “gold standard” angiogenesis assay is to be developed; thus, a combination of methods are employed to identify the effects of molecule or protein-mediated molecular/cellular signaling processes involved in angiogenesis. Under in vitro settings, two-dimensional (2D) tube formations from endothelial cells are assessed using reduced growth factor Matrigel and has offered the basis for cell-based assays concerning the evaluation of angiogenic processes. Reduced growth factor Matrigel provides a proangiogenic environment for endothelial cells. One of the greatest advantages of this method is that the whole process can be done in as short as 6–24 h depending on the cell types. This method is analyzed based on the formation of tubes (measured as tube length) in the angiogenesis process. It is rapid, robust, and adapted to assess the pro- and antiangiogenic capability of small molecules and genes targeted using small interfering RNAs (siRNAs). However, a major drawback of this method is missing the crosstalk with other cell types, which could contribute to a lack of tube stability for longer periods of time. The three-dimensional (3D) tube formation method was tailored to overcome some of these effects, but the process takes a longer (14 days) time than the 2D angiogenesis method. Over a period, ex vivo and in vivo models were developed to mimic the real-time effects of angiogenesis [5, 6]. A common approach to assess ex vivo angiogenesis is to use an explant from a specific organ/tissue and monitor the vessel outgrowth over a length of time. In this method, the explant will be covered with reduced growth factor Matrigel like a sandwich. In the current book chapter, the aortic rings are examined based on the number of sprouts and the area covered by each sprout. Aortic ring assays are widely used; however, there are some problems with variability from different animals, so using multiple rings from each animal is very important [7]. A mouse aortic ring assay can be employed to assess changes in gene function in transgenic mice [8]. One major advantage of the ex vivo method is that it mimics vascular sprouting or neovascular growth in animals. A caveat, however, is the use of large vessels which is distant from the ideal microvascular processes. In vivo angiogenesis methods were developed to overcome the difficulties associated with in vitro and ex vivo methods. Various in vivo models were established to understand microvascular angiogenic processes, including growth factor stimulated angiogenesis (implantation), tumor angiogenesis (cancer cell injection and chemically induced), oxygen-induced retinopathy (OIR), and chick embryo chorioallantoic membrane assays. Further, developmental angiogenesis was also assed using zebra fish models. Matrigel plug-based in vivo angiogenesis is assessed based

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on measuring neovascularization by staining sections of Matrigel plugs using CD31 or isolectin-B4 antibodies [9]. The prime advantage of studying angiogenesis in vivo is that it involves various cell types with the balancing of pro and antiangiogenic factors that coordinate together to control mature vessel formation. Refined reduced growth factor Matrigel is beneficial because it is extracted from mouse tumors and provides a natural environment in which an angiogenic response is initiated. Major concerns with in vivo experiments include the expense, time consumption, variation between animal models, and the presence of other growth factors and cytokines; therefore, results should be assessed with caution. This chapter mainly describes in vitro, ex vivo, and in vivo methods to assess angiogenesis by analyzing tube formation in 2D (endothelial cells plated on Matrigel) and 3D (endothelial cells mixed together with reduced growth factor Matrigel) assays, sprouting angiogenesis (aortic explants), and finally neovascularization (implanting Matrigel plugs into flanks of mice). All of these methods are routinely used in our laboratory and the results are published [10–12].

2 2.1

Materials Cell Culture

1. Mouse aortic endothelial cells (MAEC): Isolated from aortic rings and used between p2 and p5 for 2D angiogenesis assay [13]. 2. Mouse dermal microvascular endothelial cells (MDEC): Kind gift from Dr. Andrew Dudley, University of Virginia, and used between the passages of p6 and p12. Cells were cultured using mouse endothelial cell growth media [10, 14]. 3. Human microvascular endothelial cell media composition: MCDB 131 Media (1
). 4. 10% FBS. 5. Human epidermal growth factor (10 ng/mL). 6. Hydrocortisone (1–50 μg/mL stock: Initially dissolve 2 mg of hydrocortisone in 2.0 mL absolute ethanol and make it to 40 mL with sterile MCDB-131 media). 7. L-glutamine (2 mM) and 1
antibiotic mix solution. 8. Human microvascular endothelial cells (HMEC-1) were purchased from ATCC and used between the passages of p2 and p13 for 2D angiogenesis assays. Cells were cultured as previously described using MCDB-131 endothelial cell growth media [15]. 9. Ethanol (70%). 10. Sterile tips (10–200 μL; room temperature).

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11. Phosphate-buffered saline (sodium phosphate dibasic, 10.1 mM; potassium phosphate monobasic, 1.76 mM; potassium chloride, 2.7 mM; sodium chloride, 137 mM; 18.2 MΩ-cm  1 water, pH 7.4 and sterile filtered through 0.22micron filter).Trypsin-EDTA solution (0.025% of trypsin; 0.01% EDTA). 12. Conical tubes (15 and 50 mL). 13. Hemocytometer or cell counter to count the cells. 14. Trypan blue solution (0.4%). 15. Biosafety cabinet. 16. Cell culture incubator. 2.2 2D and 3D Angiogenesis

1. Matrigel: Matrigel (growth factor reduced or high concentration for in vitro; Phenol Red-free Matrigel for in vivo). To thaw Matrigel, leave it overnight at 4  C and aliquot into 1 mL tubes. Always use cold tips to handle the Matrigel. 2. Phosphate-buffered saline (PBS). 3. Ethanol (70%). 4. Sterile tips (10–200 μL; room temperature and cold stored, preferably 20  C). 5. Trypsin-EDTA solution (0.025% of trypsin; 0.01% EDTA). 6. 48-well cell culture plate. 7. Conical tubes (15 and 50 mL). 8. 1.5 mL tubes. 9. Hemocytometer or cell counter to count the cells. 10. Trypan blue solution (0.4%). 11. Microscope equipped with camera (equipped with objectives
4–
40). 12. Biosafety cabinet. 13. Cell culture incubator.

2.3 Ex Vivo Sprouting Angiogenesis

1. Mouse aorta. 2. Phosphate-buffered saline (PBS). 3. Dissecting microscope (to clean the aorta with surrounding external tissue). 4. Spring scissors to clean the aorta. 5. Dissecting plate (100 mm dish containing solidified gel/silica). 6. Silver sequin pins. 7. Growth factor reduced Matrigel. 8. 48-well cell culture plate.

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9. Sterile tips (10–200 μL; room temperature and cold stored, preferably 20  C). 10. Endothelial cell growth media. 11. Microscope (equipped with objectives
4–
40), equipped with camera to acquire images. 12. Biosafety cabinet. 13. Cell culture incubator. 14. Pointed tweezers. 15. Euthanasia according to IACUC-approved protocol. 16. Gauze. 17. Scissors to dissect the animal. 2.4 Matrigel Plug Assay Materials

1. Saline. 2. Growth factor-reduced Matrigel (Phenol Red-free; 10 mg/ mL). 3. Basic fibroblast growth factor (bFGF). 4. Vascular endothelial growth factor (VEGF). 5. Heparin sulfate/heparin. 6. Anesthesia (ketamine/xylazine cocktail according to the IACUC protocol or other approved anesthesia, euthanasia) protocols. 7. Depilated cream/hair trimmer. 8. Syringes (0.5 or 1 mL). 9. Needles (27 or 30G). 10. Mouse colony maintenance facility.

3

Methods Endothelial cells: Prior to seeding the primary endothelial cells, coat the cell culture dishes/flasks with fibronectin (5 μg/mL) or 0.1% gelatin for at least 1 h. Culture the endothelial cells at 70–90% confluence using appropriate endothelial cell growth media. Mouse endothelial cell growth media composition: low-glucose DMEM, 10% FBS, 10% Nu-Serum IV, vascular endothelial growth factor (1 ng/mL), basic fibroblast growth factor (6 ng/mL), heparin salt (0.1 mg/mL), 1% insulin-transferrin-selenium, and 1
antibiotic mix. All in vitro experiments should be conducted in the biosafety cabinet, and incubations should be kept at 37  C. Work bench area for experiments employing mice should be sterilized with 70% ethanol. Mouse aortas are extracted under sterile conditions.

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3.1 2D Angiogenesis/Tube Formation Assay

1. Prior to starting the experiment, please see Note 1. 2. Pipette out 160 μL of reduced growth factor Matrigel (10 mg/ mL) into each well of 48-well cell culture plate and keep it in the CO2 incubator for 20 min (see Note 2). 3. Gently overlay 250 μL of endothelial cell (EC) culture media on top of the Matrigel (enough to cover) (see Note 3). 4. Wash endothelial cells (70–90% confluency) with PBS and detach them with trypsin/EDTA solution (0.025%/0.01%). Once the cells are detached, neutralize the trypsin activity with endothelial cell growth media. Count the cells using a hemocytometer or automated cell counter (two to three times), and then centrifuge the cell suspension at 1200 rpm for 5 min. 5. Discard the supernatant and resuspend the cell pellet in endothelial cell growth media as 1 million cells/1 mL. 6. Aliquot the 80 μL (80,000 cells) of cell suspension into a sterile new Eppendorf tube and mix with 170 μL of endothelial cell growth media to make up the total volume to 250 μL (see Note 4). 7. Carefully remove the media from Matrigel placed well and add 250 μL of cell suspension into the wells. Incubate the plate in the cell culture incubator (see Note 5). At this point, one should optimize the number of cells required for tube formation and stability. Usually, tube formation can be noticed from 2 to 4 h with 20 to 80 K endothelial cells. Tubes from human endothelial cells last for a longer time than mouse-derived endothelial cells. Time and number of cells to form tubes vary between mouse- and human-derived cells. 8. Take bright field images (4
) every 2 h using microscope equipped with camera until tubes collapse/retract (Fig. 1). 9. Quantify tube length from the acquired images using NIH ImageJ software.

3.2 3D Angiogenesis/Tube Formation Assay

1. Prior to starting the experiment, please see Note 1. 2. Wash endothelial cells (70–90% confluency) with PBS and detach them with trypsin/EDTA solution (0.025%/0.01%). Once the cells are detached, neutralize the trypsin activity with endothelial cell growth media. Count the cells using a hemocytometer or automated cell counter (two to three times) and then centrifuge the cell suspension at 1200 rpm for 5 min. 3. Dilute the cells as 1.25 million cells per mL into a 1.5 mL tube. 4. Centrifuge the cell suspension at 1200 rpm for 5 min and discard the supernatant.

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Fig. 1 Typical images of a 2D tube formation assay for human and mouse endothelial cells. Human microvascular endothelial cells (HMEC-1) or mouse aortic endothelial cells (MAEC) (80,000/well) were plated on growth factor reduced Matrigel and tube formation was monitored, and bright field images were captured from 4, 6, and 24 h for HMEC-1 cells and 2, 4, and 8 h for MAEC cells

Fig. 2 Representative images showing formation of vessel-like structures (arrow) by mouse microvascular endothelial cells (MDEC; 1.25 million cells) in 3D Matrigel angiogenesis assay

5. Add 250 μL of Matrigel (growth factor reduced/diluted from high concentration) to the cell pellet and mix well with pipette. 6. Transfer the whole suspension into one well of a 48-well plate and incubate in the cell culture incubator for 20 min. 7. Add 300 μL of endothelial cell growth media and carefully change the media every 2 days. 8. Take bright-field images (4
) after 7, 10, and 14 days to observe the sprouting vessels—using microscope equipped with camera (Fig. 2).

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3.3 Ex Vivo Sprouting Angiogenesis Assay

1. Prior to starting the experiment, please see Notes 1 and 6. 2. Pipette out 160 μL of Matrigel into each well of 48-well plate, and incubate the plate in the cell culture incubator for 20 min. 3. Add 200–250 μL of endothelial cell growth media to Matrigel placed wells and incubate the plate in the incubator until the aortic rings are placed. 4. Euthanize the animal according to the IACUC-approved protocol, and carefully locate and start cutting along the aorta from the heart toward the abdominal aorta. Place the aorta in the dissecting plate with PBS, and using spring scissors, remove the fat tissue around the aorta. 5. Cut the aorta into 1 mm rings and keep it in 1.5 mL centrifuge tube with PBS (see Note 7). 6. Carefully remove the media from Matrigel placed wells. 7. Using pointed tweezers, carefully remove the aortic rings from the 1.5 mL centrifuge tube and place them on the Matrigelcoated wells. 8. Cover the aortic rings with an additional 160 μL of Matrigel like a sandwich. 9. Place this plate in the cell culture incubator for 20 min then add 200 μL of endothelial cell growth media into each well and return the plate into the cell culture incubator. 10. Monitor aortic explants regularly and acquire (4
) images every day starting from day 0 to day 7 or longer depending on the study. You will typically notice vessel sprouting from day 3 (Fig. 3). 11. Analyze images for the average number of sprouting vessels, which are calculated for each day, and data are expressed as fold increases/decreases depending on the experimental conditions.

Fig. 3 Ex vivo sprouting angiogenesis assay: aortic explants were extracted from WT mice (C57BL6J), sandwiched in reduced growth factor Matrigel, and sprouting angiogenesis was monitored for 7 days. Representative images (4
) showing sprouting angiogenesis from day 3 to 7

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12. Endothelial cells can be isolated or extracted from the sprouting explants using a two-step selection by anti-CD31 and antiICAM2-specific magnetic beads [16] and cultured for further use. These cells can proliferate up to 5–7 passages. 3.4 Matrigel Plug Assay

1. Prior to starting the experiment, please see Note 8. 2. Prepare Matrigel suspension by mixing 100 μL of Matrigel with bFGF (125 ng/100 μL), heparin sulphate (15 μg/100 μL), and VEGF (1.2 ng/100 μL) in a sterile 1.5 mL centrifuge tube in the biosafety cabinet, and keep it in the ice box until ready to use. 3. Anesthetize the mice using isoflurane (1.5%) or the lab-approved IACUC anesthetizing procedure and secure the mice on prone position. 4. Remove hair using depilated cream/trimmer at flank region of the mice. Sterilize the area using 70% ethanol. Using 0.5 mL syringes, collect 100 μL of premade Matrigel mix and inject subcutaneously into each side of the flank (see Note 9). 5. After injection do not take off the needle immediately; keep it in for at least 30 s. 6. Remove the mice from anesthesia and monitor them for 2 weeks. 7. Two weeks post-implantation, euthanize the mice according to the approved IACUC protocols and excise the Matrigel plugs (see Note 10). 8. Immediately fix the excised Matrigel plugs in Tissue-Tek optimum cutting temperature (OCT) compound or paraformaldehyde (4%) for histological studies. 9. Following sectioning, Matrigel plugs can be stained with CD31 or Isolectin B4 to observe the neovascularization (Fig. 4). 10. Acquire images of the vessels and analyze the microvascular density using ImageJ software.

4

Notes 1. Prior to starting the experiment, place the Matrigel in the ice bucket to thaw (at least 3 h prior to start the experiment). While pipetting the Matrigel, use only cold stored tips to release smoothly into the wells. If using 1 mL tips, cut the tip of the tips with scissors for easy handling. Keep the EC media in 37  C water bath. 2. While pipetting the Matrigel, gently release to avoid bubbles.

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Fig. 4 Matrigel plug in vivo angiogenesis assay: Matrigel mixed with VEGF and bFGF was injected into the flanks of WT mice (C57BL6/J). Matrigel plugs were extracted from mice after 14 days. Image of typical Matrigel plug. Immunofluorescence images from Matrigel plugs sections showing vessel formation, as visualized by CD31 staining. Green ¼ CD31; Blue ¼ DAPI staining for nuclei. (Adapted from Thoppil et al. [12])

3. While adding EC media, release gently dropwise to the side of the wall. 4. Adjust this total volume accordingly with the desired number of cells (20–80 K cells) to assess the tube formation. 5. Mix well before adding the cells into the well. Please make sure not to tilt the plate post-plating the cells. 6. Before isolating aorta, add 160 μL/well of growth factorreduced Matrigel into the respective labelled wells (48-well cell culture plate). Try to minimize bubble formation while adding the Matrigel into the wells. After 10 min, add 200 μL of EC media on top of the Matrigel. 7. Keep the aortic rings in ice bucket until use.

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8. This assay requires mice, preferably 6–8 weeks old (except for aging studies) for better results. Prior to starting the experiment, mix growth factorreduced Matrigel, supplemented with basic fibroblast growth factor (125 ng/plug), heparin sulfate (15 μg/plug), and vascular endothelial growth factor (1.2 ng/plug) in a 1.5 mL centrifuge tube and keep it in ice. 9. Always use cold syringes (to make syringes cold, keep the syringes in the ice for 20 min). 10. Carefully excise the Matrigel plugs; sometimes plugs could look like transparent gel. References 1. Carmeliet P, Jain RK (2011a) Molecular mechanisms and clinical applications of angiogenesis. Nature 473(7347):298–307 2. Carmeliet P, Jain RK (2011b) Principles and mechanisms of vessel normalization for cancer and other angiogenic diseases. Nat Rev Drug Discov 10(6):417–427 3. Dvorak HF, Brown LF, Detmar M, Dvorak AM (1995) Vascular permeability factor/vascular endothelial growth factor, microvascular hyperpermeability, and angiogenesis. Am J Pathol 146(5):1029–1039 4. Hoeben A, Landuyt B, Highley MS, Wildiers H, Van Oosterom AT, De Bruijn EA (2004) Vascular endothelial growth factor and angiogenesis. Pharmacol Rev 56(4):549–580. https://doi.org/10.1124/pr.56.4.3 5. Baker M, Robinson SD, Lechertier T, Barber PR, Tavora B, D’Amico G, Jones DT, Vojnovic B, Hodivala-Dilke K (2011) Use of the mouse aortic ring assay to study angiogenesis. Nat Protoc 7(1):89–104. https://doi. org/10.1038/nprot.2011.435 6. Malinda KM (2009) In vivo matrigel migration and angiogenesis assay. Methods Mol Biol 467: 287–294. https://doi.org/10.1007/978-159745-241-0_17 7. Kruger EA, Figg WD (2001) Protein binding alters the activity of suramin, carboxyamidotriazole, and UCN-01 in an ex vivo rat aortic ring angiogenesis assay. Clin Cancer Res 7(7): 1867–1872 8. Devy L, Blacher S, Grignet-Debrus C, Bajou K, Masson V, Gerard RD, Gils A, Carmeliet G, Carmeliet P, Declerck PJ, Noel A, Foidart JM (2002) The pro- or anti-

angiogenic effect of plasminogen activator inhibitor 1 is dose dependent. FASEB J 16(2):147–154. https://doi.org/10.1096/fj. 01-0552com 9. Cappelli HC, Kanugula AK, Adapala RK, Amin V, Sharma P, Midha P, Paruchuri S, Thodeti CK (2019) Mechanosensitive TRPV4 channels stabilize VE-cadherin junctions to regulate tumor vascular integrity and metastasis. Cancer Lett 442:15–20. https://doi.org/ 10.1016/j.canlet.2018.07.042 10. Adapala RK, Thoppil RJ, Ghosh K, Cappelli HC, Dudley AC, Paruchuri S, Keshamouni V, Klagsbrun M, Meszaros JG, Chilian WM, Ingber DE, Thodeti CK (2016) Activation of mechanosensitive ion channel TRPV4 normalizes tumor vasculature and improves cancer therapy. Oncogene 35(3):314–322. https:// doi.org/10.1038/onc.2015.83 11. Guarino BD, Adapala RK, Kanugula AK, Lenkey NM, Dougherty JA, Paruchuri S, Khan M, Thodeti CK (2019) Extracellular vesicles from pathological microenvironment induce endothelial cell transformation and abnormal angiogenesis via modulation of TRPV4 channels. Front Cell Dev Biol 7:344. https://doi.org/ 10.3389/fcell.2019.00344 12. Thoppil RJ, Cappelli HC, Adapala RK, Kanugula AK, Paruchuri S, Thodeti CK (2016) TRPV4 channels regulate tumor angiogenesis via modulation of Rho/Rho kinase pathway. Oncotarget 7(18):25849–25861. https://doi. org/10.18632/oncotarget.8405 13. Adapala RK, Talasila PK, Bratz IN, Zhang DX, Suzuki M, Meszaros JG, Thodeti CK (2011) PKCalpha mediates acetylcholine-induced

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activation of TRPV4-dependent calcium influx in endothelial cells. Am J Physiol Heart Circ Physiol 301(3):H757–H765 14. Dudley AC, Khan ZA, Shih SC, Kang SY, Zwaans BM, Bischoff J, Klagsbrun M (2008) Calcification of multipotent prostate tumor endothelium. Cancer Cell 14(3):201–211 15. Kanugula AK, Adapala RK, Midha P, Cappelli HC, Meszaros JG, Paruchuri S, Chilian WM, Thodeti CK (2019) Novel noncanonical regulation of soluble VEGF/VEGFR2 signaling by

mechanosensitive ion channel TRPV4. FASEB J 33(1):195–203. https://doi.org/10.1096/ fj.201800509R 16. Lim YC, Garcia-Cardena G, Allport JR, Zervoglos M, Connolly AJ, Gimbrone MA Jr, Luscinskas FW (2003) Heterogeneity of endothelial cells from different organ sites in T-cell subset recruitment. Am J Pathol 162(5): 1591–1601. https://doi.org/10.1016/ S0002-9440(10)64293-9

Chapter 10 Determination of Blood-Brain Barrier Hyperpermeability Using Intravital Microscopy O’lisa Yaa Waithe, Chinchusha Anasooya Shaji, Ed W. Childs, and Binu Tharakan Abstract The blood vessels that vascularize the central nervous system (CNS) exhibit unique properties, termed the blood-brain barrier (BBB). The BBB allows these blood vessels to tightly regulate the movement of ions, molecules, and cells between the blood and the brain. The BBB is held together by tight junctions of the neighboring endothelial cells of the barrier, more specifically by tight junction proteins (TJPs) which can take the form of either integral transmembrane proteins or accessory cytoplasmic membrane proteins. BBB permeability can furthermore be affected by various factors, including but not limited to TJP expression, size, shape, charge, and type of extravascular molecules, as well as the nature of the vascular beds. The BBB is essential for the proper maintenance of CNS function, and its structural integrity has been implicated in several disorders and conditions. For instance, it has been shown that in the cases of traumatic brain injury (TBI), TBI-associated edema, and increased intracranial pressure are primarily caused by cases of hyperpermeability seen because of BBB dysfunction. Intravital microscopy is one of the most reliable methods for measuring BBB hyperpermeability in rodent models of BBB dysfunction in vivo. Here, we describe the surgical and imaging methods to determine the changes in BBB permeability at the level of the pial microvasculature in a mouse model of TBI using intravital microscopy. Key words Blood-brain barrier, Intravital microscopy, Traumatic brain injury

1

Introduction The blood-brain barrier (BBB) is a line of defense between the cerebral capillary space and the fluids of the central nervous system. Comprised of capillary endothelial cells alongside a basement membrane, glia, and astrocytes, it serves the purpose to protect and filter. The BBB is held together by the cell-cell adhesion of the microvascular endothelial cells of the barrier [1–4]. These adhesions are maintained by tight junction proteins (TJPs), which can take the form of either integral transmembrane proteins (claudins, occludin, junctional adhesion molecules, etc.) or accessory cytoplasmic membrane proteins (e.g., Zonula occludens-1; ZO-1). It

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has been shown that knockdown of essential TJPs such as ZO-1 can increase barrier permeability, most likely via the decrease in structural integrity from the decrease in this TJP expression. Thus, it can be said that permeability of the BBB can be affected by some factors; in addition to TJP expression, barrier permeability can also be affected by the size, shape, charge, type of the extravascular molecules, as well as the nature of the vascular beds [1–4]. The BBB is a major effector in proper functionality and neurotransmission of the central nervous system, and thus its structural integrity (and thus permeability) has been brought under investigation concerning multiple disorders and conditions. In this protocol we describe impact of traumatic brain injury (TBI) on BBB hyperpermeability using intravital imaging [5–9]. For instance, it has been shown that in the cases of traumatic brain injury (TBI), TBI-associated edema, and increased intracranial pressure (ICP) were primarily caused by cases of hyperpermeability seen because of barrier damage [5–9]. Of this TBI-associated edema, vasogenic edema is shown to be the most significant form, occurring because of excessive leakage of plasma fluids and proteins from the BBB, ultimately resulting in microvascular hyperpermeability. Due to the nature of BBB permeability and its relation to disease, the use and development of reliable animal models is imperative to properly study and understand the various aspects of this barrier. One such method of accessing barrier permeability is through intravital microscopy. Intravital microscopy is a reliable method for measuring BBB permeability, done following the induction of a known effector of the BBB such as TBI injury [10–16]. Intravital imaging involves the utilization of fluorescein-isothiocyanate (FITC)tagged dextran of known molecular weight and fluorescence microscopy [4]. Ideally, one should be able to visualize the fluorescent (FITC) leakage in the BBB from the vasculature. However, before imaging, there must be a condition provoking a veritable change in the barrier. As previously mentioned, this can often be the case of TBI [8, 17–19]. Induction of TBI can be done in animal models via controlled cortical impact (CCI). One of the major benefits this method offer is the quantitative control of injury force and velocity [17, 18]. Additionally, tissue deformation and its flexibility can be easily controlled in a research lab setting to produce wide ranges of injury magnitudes while also producing gradable functional impairments, tissue damage, or both.

2 2.1

Materials Reagents

1. Alcohol (70%). 2. Saline (0.9%). 3. Phosphate-buffered saline (PBS).

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4. Betadine solution. 5. Artificial cerebrospinal fluid. 6. Trichloroacetic acid. 2.2

Equipment

1. Controlled cortical impactor (CCI devise). 2. Cooling centrifuge. 3. Homogenizer. 4. Electric shaver. 5. Micro-drill. 6. Burrs for micro-drill. 7. Timer. 8. Gaymar pump. 9. Heating pad. 10. Intravital microscope.

2.3

Supplies

1. Lubricating eye ointment. 2. Cover glass with a diameter of 5 mm. 3. Large blunt/blunt curved scissors. 4. Gloves. 5. Loctite superglue. 6. Syringes. 7. 25G, 27G/30G needles. 8. Blunt- and sharp-end scissors. 9. Blunt- and sharp-end forceps. 10. Sterile cotton-tipped applicators. 11. Gauze. 12. Standard tweezers. 13. Hemostat forceps. 14. Tray with Styrofoam cooler.

2.4 Working Solutions

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1. FITC-dextran (10 kDa). 2. Anesthetic agent—urethane (or isoflurane).

Methods

3.1 Anesthetizing the Animals

All procedures and protocols involving animals need to be approved by the relevant institutional animal care and use committee (IACUC) or relevant ethics committee. It is advised to consult with institutional veterinarian prior to the start of any animal procedures. Mice are recommended to be housed in a controlled

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environment within the target institution’s animal facility. Mice should be allowed a 1-week acclimation period if brought over from an outside institution or facility. Water and food are provided ad libitum. C57BL/6 male mice ranging from 2 to 5 months old are used for imaging experiments in this protocol (see Note 1). Anesthesia dosage is calculated based on animal weight, and thus animals’ initial weights must be taken before administration. Anesthesia type is normally based on the kind of surgery performed, the desired duration of anesthesia effect, route of administration (i.e., oral or IP), etc. (see Note 2). Injectable anesthetics are typically prepared fresh. Wakefulness of the animal should be examined prior to any procedures, typically by toe-pinch or tail-pinch response. Following anesthesia, eye lubricating ointment is applied prevent dryness of the eyes. To ensure the animal maintains proper body temperature, it must be placed under a lamp or on a heat pad throughout the duration of the procedure, as many anesthetics can cause changes in homeostatic regulation (see Notes 3 and 4). If required, a rectal probe can be used to monitor the body temperature of the animal (see Note 5). The animal should be warmed sufficiently prior to the urethane injection (via either heat pad or lamp as stated before), as well as for the duration of the procedure (see Note 6). 3.2 Intravenous Injection via Tail Vein

Intravital microscopy (IVM) utilizes the extravasation of a fluorescent probe (fluorescein-isothiocyanate-FITC-tagged compounds of known molecular weight are commonly used) for evaluating vascular permeability. The fluorescent probe is normally injected via tail vein (or jugular vein). Thus, mastering the tail vein injection technique is very important prior to any experimentation. In this method, the intravenous injection was made into the tail vein. The mouse tail has several small veins, so proper care should be taken to ensure the drug is injected into the right one (i.e., lateral tail vein). Sterility of the solutions injected as well as the workspace should be always maintained, as it can affect the animal mortality rate. If needed, dilation of the vein can be done via warming the mouse (either via heat lamp, warming pad, or immersing the tail in warm water (see Note 8)). 27/30G X ½ inch needles are used for standard tail vein injections in mice. To ensure that the drug/dye is properly heated to the body temperature of the animal, prior to injection, let it sit in a warm water bathe (around 37–38 °C). Standard tail vein injection volumes for mice do not typically exceed 200–250 μL, though the exact maximum volume may depend on the size, strain, and vehicle used. Injection of air bubbles into the animal can be lethal, and thus special care should be taken to properly prepare the dye injections sans bubbles. During the injection process, the needle containing the dye should be inserted into tail at an approximate 10–15° angle, the needle bevel facing upward. The syringe remains parallel to the tail.

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The needle should be inserted into the tail as low as possible. However, should the first injection fail, chose another secondary position slightly higher on the tail. Ideal distance of injection is 1/3 of the tail. The lack of resistance is the best determinant to successful injection into the tail vein. Excessive force and pressure are not advisable as it may apply back pressure that can lead to the collapse of the vein. Restraining devices can also be used for performing tail vein injections if needed. 3.3 Surgical Procedures

Prior to any surgical procedures, the sterility of the workspace and tools must be ensured. Sterilize surgical instruments using instrument sterilizers such as cidex. Autoclaving reused instruments in sterile sealed bags is also advised. To avoid dehydration, animals should be injected with 1 mL isotonic saline solution. Body temperature of 37–38 °C must be always maintained, via use of a heating pad or lamp. This temperature can be checked via use of a rectal probe. 1. Using the electric razor, shave the dorsal side of the animal’s head. 2. For sterility, the shaved spot should be wiped with gauze or sponge soaked in 70% ethanol, followed by betadine solution. 3. Apply the eye lubricant directly to the eyes of the animal using a sterile cotton swab. This prevents loss of moisture in the eyes (see Note 9). 4. Intravenous injection of FITC via tail vein as detailed. 5. Animals are then prepped for either sham or injury procedures for CCI. 6. Using a forceps, gently lift a small piece of skin at the midline of the head and make a small incision with the help of scissors. Remove the skin from top of the skull, exposing the sagittal suture, bregma, and lambda. 7. Apply a drop of lidocaine and epinephrine solution immediately onto the periosteum to avoid excessive bleeding or pain. 8. The periosteum underneath the skull should be gently separated by using scissors to make an incision and moving it around using sterile cotton swabs. 9. The circular craniotomy window is made using a micro-drill with a diameter ranging from 3 to 4 mm. The animal’s head should be intermittently irrigated with warmed artificial cerebrospinal fluid (ACSF) or sterile saline in order to prevent overheating of the brain, as it can hamper with the results (see Notes 10 and 11). Remove the resulting bone flap. 10. Sham animals only receive the craniotomy surgery, whereas the CCI-induced group receives both the brain injury and the following craniotomy procedure.

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11. This method uses the Benchmark stereotaxic impactor from Leica (see Note 12). Dwell time, velocity, and impactor tip size should be decided and adjusted beforehand. Setting options can be found discussed in Note 12. Impactor position is determined by the placement of the animal’s head. 3.4 Traumatic Brain Injury

1. Place the extend/retract toggle switch into the “extend” position. 2. Shams are then mounted on the stereotaxic frame. Hold the animal toward one side of the bar and introduce the other side into the ear canal and tighten after putting the fixture. 3. Center the impact probe onto the lambda and the anterior/ posterior, medial/lateral readings, and zero the digital manipulator. 4. The clip located at the end of the contact sensor’s lead wire should be pinned to the skin, ear, or tail base. Impact probe is lowered onto cranium until a beep is heard. 5. The dorsal/ventral reading on digital manipulator is zeroes, and the retract/extend toggle switch is flipped once again to the “retract” position. 6. Dorsal/ventral settings on the digital manipulator can be adjusted to negative values based on the injury depth chosen for the study. 7. Remove the animal from the CCI instrument. Prep and place it on a stereotaxic frame for imaging under the IVM. 8. Animal is then removed from the platform and placed in an incubator for recovery and monitoring prior to placement back in cage or to another pad/lamp for euthanasia.

3.5 Intravital Microscopy (IVM)

Intravital microscopy is a visualization technique used to study cellular and subcellular interactions that occur in live animals. It is hence a very useful technique for studying the various molecular and physiological changes that take place in animal tissue. Although it was initially used to study microcirculatory changes in postcapillary venules following inflammation, or in various in vivo studies employing transparent tissues like cremaster muscle, mesentery, etc., with the help of fluorescently tagged proteins or agents of known molecular weight, it can now be used to study the cell biology and dynamic functionality in opaque tissues such as the brain. The most used probes to do this are fluorescentisothiocyanate (FITC), Rhodamine 6G, and some fluorescently tagged antibodies. This method can also enable further analysis of the effect of various pharmacological agents or various genetic manipulations on the in vivo system. Conventional microscopy and IVM differ in the way the sample is handled: in vivo conditions entail a difference in the working distance between the condenser

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and the objectives as a result of the variance of thickness between the whole organism and traditional cell imaging. To compensate for the use of the in vivo animal, a large stage is needed. A larger stage is also beneficial in that it can hold a variety of additional sensors, tubing, etc. Thermostatic control of animal core and surface temperature is essential; thus, the need for a customized stage is often encountered and IVM also requires sufficient digital storage to view and record these dynamic events [4, 10, 11, 14, 16]. 3.6 Imaging Procedure

1. The surgical site is cleaned with absorbable gelatin sponges. 2. A small cover glass resembling the size and shape of the removed bone flap is placed on the surgical site. Seal the site with super glue by gently placing the glue on the outer edge of the surgical site. 3. The microscope is set up in an enclosure of warm and humidified air, or alternatively, the animal, microscope stage, and the objective must be warmed by other means (for instance, using heating pads to warm the animal and the stage while objective heaters are made available for warming up the objective). 4. The aluminum plate is inserted under the objective after being cleaned using 70% ethanol. 5. This method uses an intravital microscope with water immersion objective using a FITC imaging cube with emission wavelength of 525 nm. 6. Open the acquisition software and configure the settings of the intravital microscope as needed. To find the correct focal plane, a 4× DIC image is taken by moving the objective toward the coverslip until the microcirculation in the brain is clearly visible. 7. To visualize the fluorescence, the DIC cube is rotated to the FITC cube, and this imaging cube has an emission wavelength of 525 nm. All imaging parameters are set up on the ND acquisition tab. 8. The vessels must all be initially checked in order to evaluate the quality of the preparation. A vessel in which blood is flowing through is around 20–80 μm in diameter; this is typical for the vessels chosen for imaging in the pial microcirculation. 9. IVM can be performed every 20 min, for up to 2 h (per animal). Each spot is recorded for 30 s for video. Alternatively, a snapshot of the image is taken (see Fig. 1). 10. Baseline-integrated optical intensities should be obtained from intravascular and extravascular sites. From this, change in light intensity values should be evaluated. This intensity change can be calculated by using the given formula:

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Fig. 1 Intravital microscopic imaging of the blood-brain barrier/brain microcirculation in anesthetized mice injected with FITC-dextran as a fluorescent probe. Traumatic brain injury (TBI; or sham injury) was inflicted in mice (with or without doxycycline treatment) under anesthesia, and pial venules of 50–75 μm diameter were visualized (40×) under a Nikon intravital microscope (a). Dotted lines have been added to better differentiate the pial venules. Graphical plotting of BBB permeability as ΔI is shown below (b). BBB hyperpermeability was observed at 30, 50, and 70 min TBI versus TBI + doxycycline groups was observed ( p < 0.05). The data shown here is taken from our article (Robinson et al. [4]. Creative Commons License https://creativecommons. org/licenses/)

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ΔI = 1 - ðI i - I o Þ=I i , where I is the change in light intensity, Ii is the light intensity inside the vessel, and Io is the light intensity outside the vessel. Experimental values obtained from this calculation should be compared with the initial baseline value and expressed as percentage change. This method decreases bias between animals due to both red blood cell accumulation and changes in room lighting. 11. Following the completion of this procedure, anesthetized mice are euthanized by cervical dislocation as accepted by the IACUC.

4

Notes 1. Depending on animal age and weight, vessel permeability is subject to change. Thus, choosing close birth dates and weights is essential for accurate and valid comparison. 2. Animals can be anesthetized in several ways: either by using a combination of methods or by being individually put down via urethane (terminal) or isoflurane (4% for induction and 1.5–2% for surgery; with 40% O2 and 60% N2 via a facemask). 3. To prevent burn injuries to the animal, heating lamps should be kept at safe working distance, and heating pads should always have a cloth or gauze laid over top; animals should never be laid directly over the warm pad. Monitor the animal constantly for signs of burns. 4. Environmental conditions such as temperature and humidity are poignant factors in the animal’s stress level and thus can indirectly influence vascular permeability. These factors should be kept as similar as possible from animal to animal. 5. If using urethane as anesthetic agent, it should be gently heated in a 37 °C water bath and vortexed to dissolve any crystals. 6. If using urethane, it is used for terminal studies and special care is required for always handling it with proper PPE. Additionally, due to its hazardous nature, urethane should only be prepared and injected to animals only under a chemical hood. 7. To prevent burn injury when warming the animal’s tail, researchers must pay close attention and monitor the tail for scalding. A heat pad used should also be maintained strictly at 37–38 °C as low temperatures can lead to hypothermia, while high temperatures lead to loss of thermal regulation and death. The heat lamp should never be placed too close to the animal to prevent both burn injury and overheating.

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8. Analgesics like buprenorphine (0.01 mg/kg) or dexamethasone (0.2 mg/kg) and carprofen (5 mg/kg) can be used in consultation with a veterinarian. 9. FITC-dextran of various molecular weight can be used (10KDa is used in this experiment). 10. Appropriate probe tip among 1, 1.5, 2, and 3 mm is chosen based on the requirement of the study and the size of the animal. Travel velocity of the impactor can vary from 1.5 to 6 m/s with a travel range up to 10.1 mm and dwell time ranging from 0.1 to 9.9 s as required. 11. The animal’s head should be fixed firmly in the stereotaxic frame before impact. Changes in positioning and loose fitting can introduce variable changes in the injury level. This can be ensured by gently applying pressure on the skull to confirm placement before applying impact. 12. The numbers indicate the depth, in millimeters, into the intact dura. The impactor tip should be cleaned via wiping it gently with sterile alcohol after each impact. Further cleaning/disinfection is done via cidex after surgery. To prevent hypothermia in the animal, the stage should be heated to approx. 37–38 °C before the animal is placed. The warm pad and the animal must remain separated by soft cloth or gauze to prevent burn injury. While performing in vivo imaging, care must be taken so that the animals can tolerate surgery and avoid excessive physiological disturbances that may interfere with results. Repeat the experiment several times to ensure validity and reliability, as the technique is potentially influenced by several external factors.

Acknowledgments The authors acknowledge the support from National Institutes of Health (NIH) grant 5 SC3 NS127765-02 (BT). References 1. Alluri H, Anasooya Shaji C, Davis ML, Tharakan B (2018) A mouse controlled cortical impact model of traumatic brain injury for studying blood-brain barrier dysfunctions. In: Traumatic and ischemic injury: methods and protocols, Methods in molecular biology, vol 1717. Springer, New York, pp 37–52 2. Alluri H, Wiggins-Dohlvik K, Davis ML, Huang JH, Tharakan B (2015 Jan 28) Bloodbrain barrier dysfunction following traumatic brain injury. Metab Brain Dis 30:1093

3. Deaglio S, Robson SC (2011) Ectonucleotidases as regulators of purinergic signaling in thrombosis, inflammation, and immunity. Adv Pharmacol 61:301–332 4. Robinson BD, Isbell CL, Melge AR, Lomas AM, Shaji CA, Mohan CG, Huang JH, Tharakan B (2022) Doxycycline prevents bloodbrain barrier dysfunction and microvascular hyperpermeability after traumatic brain injury. Sci Rep 12:5415

Intravital Microscopic Imaging of the Blood-Brain Barrier in Mice 5. Gean AD, Fischbein NJ (2010) Head trauma. Neuroimaging Clin N Am 20:527–556 6. Kasper C, Yvette C (2015) Traumatic brain injury. In: Annual review of nursing research, vol 33. Springer, New York 7. Khan M, Im YB, Shunmugavel A, Gilg AG, Dhindsa RK, Singh AK, Singh IJ (2009) Administration of S- nitrosoglutathione after traumatic brain injury protects the neurovascular unit and reduces secondary injury in a rat model of controlled cortical impact. Neuroinflammation 6:32 8. Kumar P, Shen Q, Pivetti CD, Lee ES, Wu MH, Yuan SY (2009) Molecular mechanisms of endothelial hyperpermeability: implications in inflammation. Expert Rev Mol Med 11:e19 9. Maas AIR, Stocchetti N, Bullock R (2008) Moderate and severe traumatic brain injury in adults. Lancet Neurol 7:728–741 10. Marques PE, Oliveira AG, Amaral SS, NunesSilva A, Almeida AFS (2012) Intravital microscopy: taking a close look inside the living organisms. Afr J Microbiol Res 6:1603–1614 11. Masedunskas A, Porat-Shliom N, Tora M, Milberg O, Weigert R (2013) Intravital microscopy for imaging subcellular structures in live mice expressing fluorescent proteins. J Vis Exp 79:50558

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12. O’Connor WT, Smyth A, Gilchrist MD (2011) Animal models of traumatic brain injury: a critical evaluation. Pharmacol Ther 130:106–113 13. Parikh S, Koch M, Narayan RK (2007) Traumatic brain injury. Int Anesthesiol Clin 45: 119–135 14. Radu M, Chernoff J (2013) An in vivo assay to test blood vessel permeability. J Vis Exp 73: e50062 15. Shen Q, Wu MH, Yuan SY (2009) Endothelial contractile cytoskeleton and microvascular permeability. Cell Health Cytoskelet 1:43–50 16. Taqueti VR, Jaffer FA (2013) High-resolution molecular imaging via intravital microscopy: illuminating vascular biology in vivo. Integr Biol (Camb) 5:278–290 17. Unterberg AW, Stover J, Kress B, Kiening KL (2004) Edema and brain trauma. Neuroscience 129:1021–1029 18. Xiong Y, Mahmood A, Chopp M (2013) Animal models of traumatic brain injury. Nat Rev Neurosci 14:128–142 19. Yuan SY (2002) Protein kinase signaling in the modulation of microvascular permeability. Vasc Pharmacol 39:213–223

Chapter 11 Imaging and Analysis of the Dynamics of Filamentous Actin Structures in Live Endothelial Cells Jerome W. Breslin and Zeinab Y. Motawe Abstract The ability to view and record the movements of subcellular structures is a powerful tool that has accelerated the discovery and understanding of signaling mechanisms that control microvascular functions such as the control of endothelial permeability. Advances in molecular biology over the past few decades have facilitated the generation of fusion proteins in which fluorescent reporters based upon the structure of green fluorescent protein can be linked to proteins found in human endothelial cells, such as VE-cadherin or β-actin. These fusion proteins have been found to incorporate into structures alongside their native protein counterparts, allowing the dynamic visualization of how these subcellular structures are modified when cells are challenged with stimuli such as inflammatory mediators. The result of such studies has been a much more advanced view of the complex mechanisms by which endothelial cells maintain barrier properties than previously obtained by only viewing fixed cells labeled by immunofluorescence. Here, we describe our protocols that we have used to view the dynamics of actin filaments using time-lapse microscopy to record endothelial cells expressing GFP-actin and the analysis tools available to quantify dynamics of subcellular structures. Key words Endothelial cells, Actin cytoskeleton, Local lamellipodia, Junction-associated intermittent lamellipodia, Actin stress fibers

1

Introduction Visualization of the spatiotemporal nature of proteins in live endothelial cells is a key method for understanding subcellular mechanisms that determine functions such as the control of microvascular permeability. There are a variety of methods for transfection of endothelial cells using chemical agents, viral vectors, and electroporation with vectors encoding fusion proteins [1–4]. We have had reproducible success with a Nucleofector II system (Lonza, Basel Switzerland), which utilizes a combination of electroporation and optimized chemical transfection medium [5–9]. For this system it is important to be able to quickly to maximize the viability of the cells. Each transfection requires 5 × 105 cells, which after transfection are seeded onto a thin glass that allows for optimal optics for

Binu Tharakan (ed.), Vascular Hyperpermeability: Methods and Protocols, Methods in Molecular Biology, vol. 2711, https://doi.org/10.1007/978-1-0716-3429-5_11, © Springer Science+Business Media, LLC, part of Springer Nature 2024

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Fig. 1 Cultureware-chamber system for live cell imaging. (a) Closeup view of a MatTek 35-mm dish with a recessed glass coverslip in the middle of the dish, on which cells are grown for time-lapse imaging. (b) Chamber insert sold by Warner instruments that fits into the 35-mm plate and allows for superfusion of the cells with warmed medium during live cell imaging experiments

microscopic viewing at high power. Before the development of specialized cultureware for live cell imaging, the cells were seeded onto #1 glass coverslips kept in a culture dish. On the day of the imaging study, these coverslips were mounted together with a plastic top to form a chamber [7]. However, the emergence of 35-mm culture dishes with a recessed coverslip located in the center (Fig. 1a) and a corresponding perfusion insert (Fig. 1b) makes an easier and more productive alternative because the chance of accidentally breaking the coverslip during various steps of the protocol is drastically reduced. Several studies over the past decade that have used endothelial cells expressing fusion proteins have accelerated understanding of the localization of signaling proteins and their activity in response to various stimuli that affect endothelial permeability. FRET reporters have highlighted the key importance of localization of signal activation in the control of endothelial permeability. For instance, peripheral activation of RhoA stimulates enhancement of the endothelial barrier, while central activation is associated with disruption [8, 10]. In addition, fusion proteins that allow for visualization of structures that contain β-actin or VE-cadherin have helped identify small, local lamellipodia in confluent endothelial cells, also termed junctional-associated intermittent lamellipodia, as important for closing any gaps that open between endothelial cells and maintaining normal junctional VE-cadherin dynamics [4–7, 11–15]. Compared to previous approaches that relied on fixed cells labeled with dyes or fluorescent antibodies to identify subcellular structures, which only provided snapshots in time, these live cell imaging approaches have significantly expanded the horizons for understanding the complex mechanisms that control endothelial

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permeability. In this chapter, we provide the details of our transfection, time-lapse microscopic imaging, and data analysis protocols that we have developed to study dynamics of GFP-actin in live endothelial cells, which enabled discoveries such as the tight association between local lamellipodia activity and endothelial barrier integrity, and the two different actin stress fiber formation mechanisms that occur in endothelial cells in response to inflammatory stimuli [5, 6, 14, 15].

2

Materials

2.1 Transfection of Endothelial Cells with GFP-Actin (Nucleofector Method)

1. Human umbilical vein endothelial cells (HUVEC) or other cultured endothelial cells (EC). At least 500,000 cells are needed per transfection, yielding two coverslip dishes for study. 2. GFP-actin plasmid or a similar vector. 3. Nucleofector II Device (Lonza). 4. Transfection Kit (either primary EC kit or HUVEC kit from Lonza or Mirus). 5. Endothelial cell growth medium (25 mL for 1 transfection; Lonza EGM2 or EGM2-MV, or LifeLine Cell Technology VascuLife media). 6. Gelatin solution: 1.5% in 0.9% NaCl. At least 1 mL is needed per transfection. 7. Sterile PBS (at least 5 mL per transfection). 8. 0.25% Trypsin-EDTA solution (2 mL per 100-mm plate of endothelial cells to be split). 9. MatTek coverslip-bottom plates (2); part P35G-0.170-14-C. 10. Sterile microcentrifuge tubes.

2.2 Setting Up the Chamber System

1. Albumin-physiological salt solution (APSS; see Note 1) or VBM Medium (LifeLine) in a flask or bottle. 2. Vacuum grease and a cotton-tip applicator (Fisher Scientific). 3. Test agent(s). 4. Warner Instruments Dual Channel Temperature Controller (TC-344C). 5. Warner Instruments Culture Dish Incubator (DH-35iL). 6. Warner Instruments (RC-37F).

Open

Perfusion

Chamber

Insert

7. Warner Instruments Cable Assembly (Controller to Culture Dish Incubator; CC-28). 8. Warner Instruments In-Line Solution Heater (SH-27B). 9. Polyethylene tubing, PE-160 (Becton Dickinson).

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10. Peristaltic pump and tubing (Masterflex, Fisher Scientific). 11. Tubing connectors. 12. Vacuum trap assembly. 2.3 Acquisition of Data

1. Standard or confocal epifluorescent microscope system with camera, excitation light source, appropriate excitation, and emission filters for GFP and a shutter system to turn excitation light off while not obtaining images. 2. Image acquisition software that enables automated capture of time-lapse images.

2.4

3

Data Analysis

1. Image analysis software such as FIJI/ImageJ [16].

Methods Standard protocols should be followed for growing endothelial cells. For the transfection, efficiency, i.e., what percentage of the cells express a detectable amount of the protein, is paramount. A very low transfection or expression efficiency will yield too few cells for study. On the other hand, too much expression can also be a potential problem as the signal is so great that subcellular structures are not as easy to identify because of high background and potentially less specific localization of the fusion proteins being studied. In addition, when studying structures at or near junctions between cells, sometimes it is advantageous if one cell expresses the fusion protein and the other doesn’t, as a clear delineation of the border between cells becomes apparent. One final important factor pertaining to transfection is that the colocalization of the transfected fusion protein with its intended destination should be verified. In previous studies, labeling of HUVEC expressing GFP-actin with Alexa Fluor-594-phalloidin showed colocalization between the native F-actin structures and GFP-actin-containing structures, suggesting that the fusion protein indeed functioned together with its native counterpart [5]. A second area of consideration for the protocol is the microscope system to be used. Live cell imaging can be done either with conventional widefield or laser confocal fluorescence microscopy. One key factor to consider is the intensity and duration of the excitation light that will cause fluorescence of the GFP-actin in the cells. Too much exposure can cause both bleaching of signal and phototoxicity to the cells, so exposure should be limited to the minimum that can produce useful signal from the cells. Another factor of importance is the environmental system for keeping the cells in their optimal condition during the experiment. Key factors to consider are temperature stability and consistency in flow conditions. Temperature fluctuations in the chamber containing the

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cells on the microscope stage can cause changes in the focus length of between the cells and objective lens that are large enough to make images go out of focus. Measures to prevent this from happening are discussed in the notes below. After collection of time-lapse image data, several image analysis tools that are readily available in most image acquisition/analysis packages and also in the freely available NIH ImageJ software are available. Details about protocols in the build of Image J called FIJI (“FIJI is Just ImageJ”) that is freely downloadable (https://fiji.sc) are provided below [16]. Several “hot keys” on the computer keyboard that help ease the workflow in ImageJ are provided in the details of the analysis portion of the protocol. 3.1 Transfection of Endothelial Cells with GFP-Actin (Nucleofector Method)

1. This protocol outlines the procedures for transfection of HUVEC using the Nucleofector II device. Some modifications may be needed to optimize transfection of other endothelial cell types. 2. Aliquot the PBS, Trypsin-EDTA, and VGM-MV medium and place in the 37 °C water or bead bath. Warm the gelatin solution as well. 3. Get the Nucleofector solution (primary endothelial or HUVEC kit) from the refrigerator and place in the biosafety cabinet to allow it to warm to room temperature. If the solution needs to be prepared, see the Nucleofector instructions. 4. Place a Nucleofector cuvette (1 per transfection) and Nucleofector transfer pipette (1 per transfection) in the biosafety cabinet. Remove the cuvette from its packaging, and open the edge of the transfer pipette packaging so it can be quickly removed later. 5. In the MatTek 35-mm coverslip-bottom plates (two per transfection), coat the recessed coverslip with gelatin solution. This only requires 200 μL of the gelatin solution per coverslip. 6. In one of the sterile microcentrifuge tubes, place 300 μL of VBM-MV. Close the cap, and place in the 37 °C, 5% CO2 incubator. 7. Turn on the Nucleofector II and select program A-34 for HUVEC, or the optimal program determined for your endothelial cell type. 8. Aspirate media from the plate of cells to be split. 9. Add warm PBS (1 mL for a 60-mm plate, or 2–3 mL for a 100-mm plate) and aspirate. 10. Add warm trypsin-EDTA (1 mL for a 60-mm plate, or 1–2 mL for a 100-mm plate). 11. Allow the cells to detach. The plate can be rapped on a counter to facilitate detachment. Confirm by microscopic observation.

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12. Add warm VBM-MV to the plate at a volume equal to the trypsin-EDTA solution that was added in step 3, and then remove this cell suspension and place in a sterile 15-mL conical tube. 13. Take a 10-μL aliquot of the cell suspension for cell counting using a hematocytometer. 14. After the counting is done, determine the volume of cell suspension that is needed to obtain 500,000 cells (for one transfection). Spin the volume that you need for the transfections for 3 min at 300×g. 15. While waiting, get the GFP-actin plasmid solution from the refrigerator and place in the biosafety cabinet. 16. After centrifugation, bring the conical tube(s) containing the pelleted cells back to the biosafety cabinet. When aspirating the medium, tilt the conical tube sideways to get as much off as possible without disturbing the pellet. 17. Resuspend the pellet in 100 μL of Nucleofector solution (see Note 2). 18. Add GFP-actin plasmid (0.2–2.0 μg) to the cell suspension. 2 μg will generate high expression, while 0.2 μg will generate a lower, less intense expression. 19. With a 1000 μL micropipette, gently mix up and down once, and then collect the 100+μL of cell suspension and place in a Nucleofector cuvette. 20. Cap the cuvette, tap down a few times, and place in the Nucleofector II device. Press start to run the program. After the program has run, the display will indicate “OK.” If an error occurs, tap the cuvette down a few more times and try again. Errors occur when the solution is not in contact with both electrodes in the cuvette. 21. Bring the cuvette back to the biosafety cabinet. Get the microcentrifuge tube containing 300 μL of warm VBM-MV medium from the incubator. Uncap the cuvette. With the sterile transfer pipette that comes with the kit, place the 300 μL of VBM-MV into the cuvette, and without mixing up and down, gently remove all of the contents of the cuvette and place them back into the microcentrifuge tube. Then place the tube in the incubator and wait 10 min before seeding the cells from this suspension (see Note 3). 22. Seed 200 μL of the transfected cell suspension onto each of the MatTek plates, only covering the recessed coverglass. Limiting the seeding to this area will make the cells form a confluent monolayer quickly.

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23. Carefully place the MatTek plates into the incubator. 24. Wait at least 1 h (up to 4 h). Then, add 1.5 mL of VBM-MV medium to each plate and incubate overnight. 3.2 Setting Up the Chamber System

1. The expression of GFP-actin is generally optimal in the first 24 h after transfection using the Nucleofector II system. The cells can be used up to 48 h after transfection; however, because of the transient nature of the transfection, after 48 h the fluorescent signal starts to be noticeable in vesicular structures due to turnover of the protein. 2. Before starting the experiment, observe the cells using a fluorescent microscope to check that they are confluent and expressing GFP-actin. If they are ready for the experiment, return the cells to the incubator for the time being as setup begins. 3. Fluid is fed through the system using a peristaltic pump. For our system, we run at a flow rate of ~1 mL/min. Add an appropriate amount of APSS to flask that will serve as the reservoir taking into consideration the anticipated length of time the experiment will last. Insert the inlet tubing that leads to the pump into the flask, and be sure that the tubing exiting the pump is connected to the In-Line Solution Heater. Run the pump to introduce fluid into the line. You can let some excess fluid drip out of the In-Line Solution Heater into a flask. 4. Apply a thin layer of vacuum grease to the bottom of the Open Perfusion Chamber Insert using a cotton tip applicator (Fig. 2a). 5. Bring one of the MatTek dishes containing cells to the microscope system, and remove the media in the dish using a 1000 μL pipette, saving ~1 mL of medium in the pipette tip (do not discard). 6. Press the Open Perfusion Chamber Insert into the dish to make a tight seal. 7. Gently dispense the media that is still in the 1000 μL pipette’s tip into the chamber to prevent the cells from drying out. 8. Place the MatTek dish with chamber insert into the Culture Dish Incubator and secure with the two clips. Attach the inlet tubing to the In-Line Solution Heater, and attach the outflow to a vacuum line (Fig. 2b). 9. Turn on the Warner Dual Channel Temperature Controller, and make sure the individual switches for the Culture Dish Incubator and In-Line Solution Heater are both on. 10. Turn on the Peristaltic Pump so that fluid enters into the chamber, and at the same time, double-check that the vacuum line is removing excess fluid, but not too much fluid from the chamber. The height of the vacuum may need to be adjusted

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Fig. 2 Setting up the chamber for live cell imaging. (a) Vacuum grease is applied to the bottom of the chamber insert in order to make a tight seal with the MatTek dish. (b) After placing the chamber insert into the dish and putting media on the cells, the dish is placed in the culture dish incubator, and an inlet line is attached to the inline heater unit (right) while a vacuum line is attached to the vacuum assembly in the dish. (c) The magnetic cover of the culture dish incubator is attached to help prevent heat loss. A glass coverslip is placed on the opening on top. In this image, the coverslip is placed partially to the side to allow micropipette access to the cells for rapid delivery of test compounds. These can also be added by wash-in via the inlet tubing if desired

slightly. When finished, the magnetic cover for the Culture Dish Incubator may be placed on top. A cover glass may also be placed on the magnetic cover to prevent heat loss. We place it partially covering the opening to be able to pipette test compounds onto the cells (Fig. 2c). 11. The system should be given about 30 min to warm up to a stable 36–38 °C steady-state temperature. Continue to monitor temperature and fluid flow throughout the experiment. 12. Turn on all of the components of the microscope system and start the acquisition software. 3.3 Acquisition of Data

1. Focus on the cells, select the desired magnification (see Note 4), and find a location for study. 2. Check or optimize the acquisition parameters for the experiment. Consider how many images will be taken over the time course and the potential for fading of signal due to excess excitation light. On our microscope system (see Note 5), which uses a 300 W xenon lamp excitation source, an exposure time of 0.5–1.0 s typically can allow for collecting 480 images over 2 h, causing a noticeable fade over time, but still allowing enough signal to get crisp images. This will vary between different microscope systems (see Note 6). Many cameras offer a binning option in which pixels can be joined together to increase sensitivity of the camera (see Note 7). 3. Set the software to collect images at the desired frequency and duration. For experiments to study the dynamics of structures

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containing GFP-actin in cells at 37 °C, four frames per minute is sufficient to capture most cell movements (see Note 8). 4. If your software allows for streaming of images to disk, this option should be used in case of any problems with power or software issues during the experiment. 5. During experiments, it is possible that the cells may drift out of focus if the temperature is not stable (see Note 9). This can be alleviated by making sure the temperature remains within a degree of the set point (37 °C) throughout the experiment. Most software packages allow for focusing between capture of images, if needed. 6. A typical protocol is to have a 30-min baseline, and then add a test agent and monitor the cells for up to 2 h. 7. Save the image file at the end of the experiment for offline analysis. 8. Turn off all equipment and clean the chamber insert after use, and flush all tubing with water. The vacuum trap will need to be cleared from time to time. Its contents should be mixed with 10% bleach before pouring down the drain. 3.4 Analysis of Imaging Data

1. Multiple software packages have the features to do the various analyses presented here. A convenient and free software package we use is FIJI, which can be installed on a Windows or Mac laptop computer so analysis can be performed offline, anywhere. 2. Images should be saved a TIF stack (single file with all images) for work in FIJI. Often, FIJI can open various file formats from different software platforms, but once the file has been opened in FIJI, it is easiest to save it as a TIF for future work. If multiple channels were used, a multichannel TIF format should be used. 3. Keep notes from the metadata about the image acquisition particularly the interval between images, μm per pixel, and time points at which any test compounds were added. 4. When opening a stack file in FIJI, note that in the upper left corner of the image window, the slice number and total number of slices will be displayed, along with the filename, number of pixels in the x and y directions (or x-y distances in μm), file type, and size. The scrollbar on the bottom controls movement between slices. If a multichannel image stack is opened, a second scrollbar for the channels is also present. When hovering the pointer over the image, the x, y, and z pixel information and pixel intensity are displayed in the bottom of the ImageJ toolbar. 5. For simplicity, the analyses discussed below are for images collected in a single plane. If z-stacks are collected at each

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time point, z-projections of images at each time point could be used. The following parameters to be analyzed below will be discussed: (a) Frequency of local lamellipodia protrusions. (b) Protrusion distance, time, and velocity. (c) Withdrawal distance, time, and velocity. (d) Actin stress fiber quantification and movement over time. 6. Choose cells for study based upon predefined inclusion and exclusion criteria (see Note 10). For studies of lamellipodia, it is easiest to choose cells expressing GFP-actin that are adjacent to cells not expressing GFP-actin. 7. Protrusion frequency: The basic protocol is to divide the cell membrane into small areas that can be observed and then scrolling through the image set, and making a note of when each local lamellipodium starts to protrude. This can be marked in the image set using the region of interest (ROI) feature in FIJI. In the FIJI menu bar, select the point tool, which can be used to mark the site of the appearance of a lamellipodium. After marking the point, press the “T” key on the keyboard, which is the hot key for marking the spot as an ROI. The ROI Manager will open when marking a new ROI. Checking the “Show All” box will allow for all ROIs to be visible, but they will only be visible in the image of the z-stack in which they are marked. All of the lamellipodia for the cell can be marked in this way over time. When selecting all of the ROIs, and then choosing “Measure” in the ROI Manager window, a “Results” window opens which lists the data for each ROI, which should include the slice number in which it appears (see Note 11). This output can be saved or copied to Excel. Each slice number will correspond to a time point in the time lapse, and the appearance of lamellipodia can be mapped to time intervals defined by the user (e.g., number per 5-min interval). For large datasets, when counting these manually would be cumbersome, the histogram function in Excel can be used to calculate these numbers. 8. Because every cell has its own unique perimeter, normalization of protrusion frequency to cell perimeter should be done to compare different cells. A rough perimeter should be drawn using the Polygon Selection tool in FIJI. Small lamellipodia do not need to be taken into account, just a more general perimeter. This can be done at the beginning and end of the time course studied, and averaged (perimeter should be selected in the Set Measurements window; see Note 11). Then the number of lamellipodia can be divided by the entire perimeter to get the number per μm. Multiplying this number by 100 generally

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Fig. 3 Example of preselected areas of the edge of a cell for the study of local lamellipodia protrusion and withdrawal dynamics. (a) Scheme in which 12 lines are drawn like an analog clock. (b) Scheme with eight lines drawn. Using preselected schemes helps reduce bias in the sampling of lamellipodia

gives number that range from 1 to 10 (number of lamellipodia per 100 μm per time interval).

9. Generating a kymograph: To obtain additional data about protrusion and withdrawal of lamellipodia, the straight line tool should be selected in the ImageJ menu. For this analysis, an unbiased way to perform the analysis is to draw 12 lines that are perpendicular to the edge of the cell in at 12, 1, 2, 3 o’clock, etc. (Fig. 3a). A smaller number of lines, such as eight lines (Fig. 3b) would also be acceptable. The key is to have enough lines to get a good sampling. After each line is drawn, press the “T” key to mark it as a ROI. After all the lines are drawn, the ROI file should be saved. Next, select the first ROI from the ROI menu (Fig. 4a), and then press the “/” key, which will bring up a window called “Reslice” that is used to draw kymographs. Only the “Avoid interpolation” box should be checked. Click “OK” and a new line-scan (kymograph) image will appear in which the y-axis represents the distance of the ROI line selected and the x-axis is time (Fig. 4b; see Note 12). 10. Protrusion distance, time, and velocity can be obtained by locating the protrusions on the kymograph and drawing straight lines along the edges where they are apparent (Fig. 4c). After drawing these lines and saving them as ROIs, on the menu bar go to “Analyze” and “Set Measurements” and make sure that “Bounding Rectangle” is checked. Then select all of the ROIs and press “Measure” and a “Results” window will appear. The BX and BY values provide information about the origin of the line. The width provides the length of the bounding rectangle and when multiplied by the time interval represents the protrusion time. The height represents the distance that the lamellipodium traveled and is the protrusion distance. Dividing the

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Fig. 4 Analysis of lamellipodia using the kymograph function in FIJI. (a) A kymograph (also called line scan) is generated by drawing a line in the image, in this case perpendicular to the edge of a cell, and then pressing the “/” key in FIJI. (b) The resulting output is a kymograph drawn by scanning the intensities of pixels along input line through the whole time-lapse image set. In the kymograph shown, the x-axis represents time and the y-axis is distance. (c) To study lamellipodia, they protrusions of the membrane are identified in the kymograph and denoted by line ROIs, and the bounding rectangles of these lines are determined using the Analyze>Measure function. (d) From these measurements, the protrusion velocity, distance, and persistence corresponding to each lamellipodium are calculated as shown. (e) Lines are also drawn over the periods during lamellipodia withdrawal, and the withdrawal distance, velocity, and time are determined in a similar fashion. Reproduced from Breslin et al. [5], with permission

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Fig. 5 Actin stress fiber formation and lateral movements, assessed by kymograph analysis. (a). In a timelapse image set of endothelial cells that are expressing GFP-actin, a line is drawn across the center of a single cell that is roughly perpendicular to any long-actin cables is drawn in order to generate a kymograph. (b) In the resulting kymograph, the x-axis represents distance, and the y-axis represents time. The brighter pixels that form lines horizontally represent where stress fibers crossed the plane of the line drawn in panel (b) Analysis of these structures is performed by superimposing user-drawn lines over these brighter areas (c). The Analyze>Measure function in FIJI is used to obtain the bounding rectangle, from which lateral velocity of actin stress fibers can be determined. In addition, the number of actin stress fibers is determined simply by the number of lines observed in a particular time frame. (Reproduced from Breslin et al. [5], with permission)

protrusion distance by the protrusion time provides the protrusion velocity for each lamellipodium (Fig. 4d). 11. Withdrawal distance, time, and velocity can be obtained by drawing lines on the lamellipodia as they regress back toward the cell center (Fig. 4e). The same methods as in step 10 above for protrusions are used to obtain the withdrawal distance, time, and velocity, just in the reverse direction. 12. Actin stress fiber number and velocity can both be determined using kymograph analysis. The stress fibers generally align in the same general direction, so that a single kymograph line that is roughly perpendicular to the actin stress fibers can be drawn across the center of the cell to produce a kymograph (Fig. 5a). The resulting kymograph will provide information both about the number and movement of actin stress fibers (Fig. 5b). It is important to consider that there are different types of actin stress fibers and multiple mechanisms by which actin stress fibers can form [17, 18]. Ventral stress fibers tend to assemble from very small filaments in central areas of the cells, while other fibers that resemble the transverse arcs observed at the leading edges of migrating cells form from cortical actin fibers at the periphery and sometimes divide laterally into two parallel stress fibers [5]. In addition, the transverse arc-like stress fibers tend to migrate toward the cell center and eventually

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disassemble [5]. These events typically can be all captured in the kymographs. Actin stress fiber number can be counted at each selected time point to be studied. The lateral movement of actin fibers, which we call actin stress fiber velocity can be determined in the same fashion as for lamellipodia, by drawing a line over the stress fiber’s movement from time point A to time point B during a particular time period (Fig. 5c). Our convention in our previous papers [5, 15] is that when a stress fiber moves toward the cell center, we assign a positive velocity, while when one moves toward the cell periphery, we assign a negative velocity.

4

Notes 1. The albumin physiological salt solution (APSS) was originally designed for tissue perfusion but is adequate for perfusion of cells for a few hours. When several hundred mL are used for an experiment, APSS is a significantly less expensive option than purchasing cell culture media. The contents of APSS are 120 mM NaCl, 4.7 mM KCl, 2 mM CaCl2·2H2O, 1.2 mM MgSO4·7H2O, 1.2 mM NaH2PO4, 2 mM Na pyruvate, 5 mM glucose, 0.02 mM EDTA, 3 mM MOPS, and 1% BSA. It can be made using the formulas provided for a Ringer’s stock solution in Table 1 and APSS in Table 2. It should be sterile-filtered in the biosafety cabinet and can be stored in aliquots at 4 °C. It is recommended that the APSS be used within 3 months when stored at 4 °C. 2. The Nucleofector solution is toxic to the cells, so it is important to work quickly during the steps when the cells are suspended in this solution. 3. The placement of this cell suspension into growth medium at this stage will help the cells recover after the electroporation and will help minimize cell death from the shock of the transfection. Table 1 Composition for 1 L of 5× Ringer stock solution Compound

Grams

mM

NaCl

35

600

KCl

1.75

23.5

CaCl2
H2O

1.47

10

MgSO4
7H2O

1.44

5.8

Bring to volume of 1 L with distilled water

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Table 2 Composition for 1 L APSS Compound

Amount

Ringer 5× stock

200 mL

mM

NaCl

120

KCl

4.7

CaCl2

2

MgSO4

1.17

0.6 M MOPS buffer (in H2O)

5 mL

3

NaH2PO4
H2O

0.168 g

1.2

Na pyruvate

0.22 g

2

Na EDTA

0.0074 g

0.02

Bovine albumin

10 g

(1%)

Glucose

0.901 g

5.1

Bring to volume of 1 L with distilled water

4. The degree of magnification will depend upon the study aims. A 100× objective with a high numerical aperture provides the best spatial resolution and transmission of signal but also only allows view of a small number of endothelial cells. 40× and 60× objectives are often good for getting data from up to 10–20 cells at a time, depending on the expression efficiency of the GFP-actin. The use of a 10× objective is fine for viewing many cells at a time, but subcellular signals are much harder to view because at 10× there are several μm per pixel so some details are not evident at this magnification. 5. Our microscope system is a custom-built Applied Scientific Imaging Rapid Automated Modular Microscope system, with an automated XY stage, automated focus, automated objective changer with Olympus Plan Fluor objectives (10×, 40×, and 100×), a Sutter Lambda LS 300 W xenon lamp for excitation light with a Lambda 10–3 controller and filter wheel, a Chroma Scientific excitation filter set and bandpass emission filter for imaging in four channels (UV, green, red, far red), and a Photometrics HQ2 camera. The system is run by a PC with ImageJ-based Micro-Manager software. 6. The shortest duration of exposure to the excitation light should be used to minimize potential phototoxicity to the cells, which can alter results. 7. Binning set at 1 × 1 will provide the best resolution, but in cases we have set binning to 2 × 2 to reduce exposure times to the

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excitation light. While this reduces precision of the measurements of structures observed in the cells, the tradeoff has less phototoxicity. 8. It is always a good idea to take a brightfield image in addition to the fluorescent images if the cells are confluent, but not all of the cells are expressing GFP-actin or the fusion protein of choice. Some microscope systems also offer the option of recording a brightfield image between each fluorescent image in the time lapse, and the two can later be overlaid. 9. It is important to monitor the experiment as images are acquired, because sometimes focus drift can occur when there are small changes in temperature within the chamber. In this case, perform a quick refocus to avoid altering the time interval between images. This can be done via the eyepieces or via the screen if the software has a refocus option. In some instances, a very fine adjustment of the focus can be made blindly, guessing which direction to focus, with a 50% chance of improving the focus, and if wrong, it can be corrected in the next interval between frames. The best way to prevent the images from going out of focus during the experiment is to optimize the inflow and vacuum lines so to get steady flow over the cells, using the same protocol for all experiments. In some instances, traffic of people into and out of the microscope room can also cause problems if there is a noticeable temperature change or draft whenever a door opens and closes. Drafts from ceiling air vents can also be problematic, and if the microscope must be placed under an air vent, installing a shield to redirect the drafts away from the microscope will help. There is also the alternative to use a plexiglass incubator chamber on the stage that keeps the cells and objective all contained in a 37 °C with 5% CO2 environment. A nice feature of this approach is that cells can be left in standard medium overnight if desired, without the need for flowing medium over the plate. However, a drawback is that access to the cells is limited, as doors need to be opened to be able to pipette test compounds on to the cells. In addition, the dish cover needs to be removed, which can easily result in accidentally moving the dish and forever losing the opportunity to see how the cells that were being observed prior to pipetting changed their behavior after the test compound was added. 10. Some cells may not be suitable for study. For instance, cells that divide during the experiment may not be useful because of the dramatic cytoskeletal changes that are attributable to cell division which may mask the changes observed in nondividing cells that are caused by the conditions being tested. Additionally, one inclusion criterion may be that the entire cell should be visible for the entire time-lapse image set. If a cell migrates

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partially out of the field of view, it might not be so useful for study. 11. If the slice number does not appear in the “Results,” this is because it was not selected as one of the data points to be measured. This can be fixed by selecting the “Analyze” dropdown menu, and then “Set Measurements. . .” which opens a dialog box with the different parameters available. Check “Stack position” to get the slice number of each ROI. 12. For the kymographs, the y-axis will represent distance either in μm or pixels depending upon the input image that was used, and the x-axis represents time. The units in the upper left corner of the image might say “μm,” but this is actually only for the y-axis. Going to the menu bar and selecting “Image” and then “Properties” will show the assignment of μm per pixels, and typically the kymograph output converts this to 1.0 for the x-axis and retains the value from the input image for the y-axis. The time interval can be assigned to the x-axis values for the analysis.

Acknowledgments While preparing this manuscript, Dr. Breslin was supported by NIH Grant R01GM120774, and Dr. Motawe was supported by an Edith Wright Hartley Graduate Scholarship. References 1. Hu YL, Chien S (2007) Dynamic motion of paxillin on actin filaments in living endothelial cells. Biochem Biophys Res Commun 357: 871–876 2. Herron CR, Lowery AM, Hollister PR et al (2011) p120 regulates endothelial permeability independently of its NH2 terminus and rho binding. Am J Physiol Heart Circ Physiol 300: H36–H48 3. Gaudreault N, Perrin RM, Guo M et al (2008) Counter regulatory effects of PKCbetaII and PKCdelta on coronary endothelial permeability. Arterioscler Thromb Vasc Biol 28:1527– 1533 4. Martinelli R, Kamei M, Sage PT et al (2013) Release of cellular tension signals selfrestorative ventral lamellipodia to heal barrier micro-wounds. J Cell Biol 201:449–465 5. Breslin JW, Zhang XE, Worthylake RA et al (2015) Involvement of local lamellipodia in endothelial barrier function. PLoS One 10: e0117970

6. Breslin JW, Daines DA, Doggett TM et al (2016) Rnd3 as a novel target to ameliorate microvascular leakage. J Am Heart Assoc 5: e003336 7. Doggett TM, Breslin JW (2011) Study of the actin cytoskeleton in live endothelial cells expressing GFP-actin. J Vis Exp 57:3187 8. Zhang XE, Adderley SP, Breslin JW (2016) Activation of RhoA, but not Rac1, mediates early stages of S1P-induced endothelial barrier enhancement. PLoS One 11:e0155490 9. Doggett TM, Breslin JW (2014) Acute alcohol intoxication-induced microvascular leakage. Alcohol Clin Exp Res 38:2414–2426 10. Szulcek R, Beckers CM, Hodzic J et al (2013) Localized RhoA GTPase activity regulates dynamics of endothelial monolayer integrity. Cardiovasc Res 99:471–482 11. Abu Taha A, Schnittler HJ (2014) Dynamics between actin and the VE-cadherin/catenin complex: novel aspects of the ARP2/3 complex in regulation of endothelial junctions. Cell Adhes Migr 8:125–135

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12. Abu Taha A, Taha M, Seebach J et al (2014) ARP2/3-mediated junction-associated lamellipodia control VE-cadherin-based cell junction dynamics and maintain monolayer integrity. Mol Biol Cell 25:245–256 13. Seebach J, Taha AA, Lenk J et al (2015) The CellBorderTracker, a novel tool to quantitatively analyze spatiotemporal endothelial junction dynamics at the subcellular level. Histochem Cell Biol 144:517–532 14. Alves NG, Motawe ZY, Yuan SY et al (2018) Endothelial protrusions in junctional integrity and barrier function. Curr Top Membr 82:93– 140 15. Adderley SP, Lawrence C, Madonia E et al (2015) Histamine activates p38 MAP kinase

and alters local lamellipodia dynamics, reducing endothelial barrier integrity and eliciting central movement of actin fibers. Am J Physiol Cell Physiol 309:C51–C59 16. Schindelin J, Arganda-Carreras I, Frise E et al (2012) Fiji: an open-source platform for biological-image analysis. Nat Methods 9: 676–682 17. Hotulainen P, Lappalainen P (2006) Stress fibers are generated by two distinct actin assembly mechanisms in motile cells. J Cell Biol 173: 383–394 18. Pellegrin S, Mellor H (2007) Actin stress fibres. J Cell Sci 120:3491–3499

Chapter 12 Isolation and Culture of Human Umbilical Vein Endothelial Cells (HUVECs) Shivam Chandel, Rathnakumar Kumaragurubaran, Hemant Giri, and Madhulika Dixit Abstract This protocol describes a simple and an economical method for isolation of endothelial cells from human umbilical vein. Umbilical cord is easily available postpartum following informed consent, and the method for its collection is noninvasive with few ethical concerns. Thus, umbilical vein is an ideal source for isolation of endothelial cells of human origin. Endothelial cells are isolated from umbilical vein by collagenase digestion. This is followed by their culture on extracellular matrix (ECM) protein coated tissue culture flasks in presence of endothelial cell growth medium. In the last few decades, human umbilical vein endothelial cells (HUVECs) have come to be regarded as a standard model system by vascular biologists to understand general principles of endothelial cell biology and dysfunction. Here, we describe isolation, culture, as well as validation of HUVECs. Key words Primary culture, Human umbilical vein endothelial cells (HUVECs), Endothelial cell isolation

1

Introduction Endothelium constitutes the innermost lining of blood and lymphatic vessels (Fig. 1a), where it plays an indispensable role in vascular homeostasis. In healthy physiological state, endothelial cells are quiescent with a low proliferative index, and they ensure that the inner vascular surface remains anti-thrombotic and antiinflammatory [1–3]. They also facilitate flow-mediated endothelium-dependent vasodilation [4]. The interface formed by the endothelium between the circulating blood and the underlying tissue is the primary site for exchange of nutrients and gases in the capillary bed of the tissues. Additionally, it controls the regulated movement of leukocytes in and out of the blood vessels (diapedesis) during the process of tissue inflammation and repair. Thus, thanks to their anatomical location, endothelial cells are the first responders to any physiological or pathophysiological changes that

Binu Tharakan (ed.), Vascular Hyperpermeability: Methods and Protocols, Methods in Molecular Biology, vol. 2711, https://doi.org/10.1007/978-1-0716-3429-5_12, © Springer Science+Business Media, LLC, part of Springer Nature 2024

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A

B Umbilical arteries

Tunica media

(Smooth muscle cells)

Tunica externa

Endothelium Internal elastic membrane

Tunica intima

Umbilical vein

C 1. Surgical grade scissor 2. 12 ml disposable syringe 3. Forcep 4. Hydrostat clamp 5. Cord clamp 6. Butterfly needle 7. Angiocath

Fig. 1 (a) Schematic representation of different layers of a blood vessel. (b) Schematic representation of a cross section of an umbilical cord depicting umbilical arteries and umbilical vein. (c) Set of surgical instruments required for the isolation protocol

occur in the biochemistry or rheology of blood. Undesired perturbations in the endothelium also referred to as “endothelial dysfunction,” in fact, is the triggering point for initiation and progression of multiple cardiovascular diseases such as hypertension, coronary artery disease, stroke, as well as tumor progression [5–7]. Endothelial cells also promote angiogenesis and formation of collateral vessels to overcome tissue ischemia due to atheromatous arterial blocks [2, 8]. Hence, cultured primary endothelial cells have been widely employed by researchers as a suitable in vitro model system to understand the biology of endothelial cells and to elucidate their role in pathogenesis of vascular diseases. Primary endothelial cells can be obtained from several animal sources, such as bovine and porcine aorta or from mice lungs and brain [9–11]. However, obtaining endothelial cells from human blood vessels such as the femoral, coronary, or the pulmonary artery has been challenging. Thankfully, easy availability of umbilical cords postpartum, has facilitated the standardization of a simple and economical procedure for isolation and culture of a relatively pure population of human umbilical vein derived endothelial cells

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(HUVECs) [12–14]. Despite their fetal and venous origin, over the last two decades, HUVECs have become the preferred cell culture model system for general understanding of endothelial biology and its dysfunction. Multiple studies by employing HUVECs, have provided critical insights on the role of endothelial cells in the biology of angiogenesis, atherosclerosis, and nitric oxide-mediated endothelial responses [15–17]. In this chapter, we provide a detailed protocol for isolation, culture, and validation of HUVECs from umbilical cord.

2

Materials

2.1 Reagents for Isolation and Culture

1. Cell culture compatible human plasma-derived lyophilized fibronectin. 2. Lyophilized collagenase (type IA) powder. 3. Phosphate-buffered saline (PBS). 4. EGM™ BulletKit™ medium (Lonza, CC3124). 5. Fetal bovine serum. 6. Cell culture-compatible penicillin-streptomycin solution. 7. Gentamicin solution. 8. 0.25% trypsin-0.02% EDTA in Dulbecco’s phosphate-buffered saline. 9. Solubilized endotoxin-free basement membrane preparation (Corning® Matrigel®, 356230).

2.2 Reagents for Characterization and Validation

1. VE-cadherin antibody (Santa Cruz Biotechnology sc-9989). 2. NOS3 (eNOS) antibody (Santa Cruz Biotechnology sc-654). 3. von Willebrand Factor (vWF) antibody (Santa Cruz Biotechnology sc-53466). 4. Alexa 594 secondary anti mouse (Invitrogen A21203). 5. Alexa 488 secondary anti rabbit (Invitrogen A32731). 6. Dil-acetylated LDL (Invitrogen, L3484). 7. Ulex europaeus-derived Agglutinin I (Ulex-Lectin).

2.3 Instrument and Other Materials

1. 12 mL syringes. 2. Dissection kit (sterile latex gloves, surgical-grade forceps, scissors, 21-gauge butterfly needles, hydrostat clamps, and cord clamps). 3. Cell culture hood with vertical laminar air flow (biosafety level class II A/B). 4. CO2 incubator with temperature control. It is preferred to have a water jacketed incubator for better temperature control.

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5. Water bath with temperature control. 6. Centrifuge with swinging bucket rotor. 7. Inverted immunofluorescence microscope with phase contrast functionality. 2.4 Preparation of Reagents and Solutions

It is preferable to use autoclaved deionized distilled water for preparation of reagents and buffers unless stated otherwise.

2.4.1 Sample Collection Buffer

Umbilical cords should be collected in 100 mL of 1X phosphatebuffered saline (PBS). The composition of PBS is 137 mM NaCl, 2.7 mM KCl, 10 mM Na2HPO4, 1.8 mM KH2PO4, pH 7.4. Add penicillin-streptomycin and gentamicin solutions to 100 mL of 1X PBS to get working concentrations of 50 U/mL penicillin, 50 μg/ mL streptomycin, and 50 μg/mL gentamicin. Store the sample collection buffer at 4°C. Prepare fresh buffer for cord collection. Sample collection buffer can be stored up to 5 days at 4 °C.

2.4.2 Fibronectin Solution for Coating

Fibronectin preparation, coating, and storage must be performed under sterile conditions in a laminar flow hood. Dissolve 1 mg of fibronectin in 32 mL of sterile Milli-Q water to get a final working concentration of ~30 μg/mL. Use 500 μL of fibronectin to coat one T-25 flask. Following addition of fibronectin solution, gently swirl the culture flask to fully cover the bottom surface of the culture flask. Incubate for 2 h at room temperature in cell culture hood under sterile conditions. Aspirate excess fibronectin solution using sterile pipette tip. Fibronectin-coated flasks can be stored at 4°C for up to 2–4 weeks, and the sterile fibronectin solution can be stored at 4°C for 2 months. Collagen or gelatin can also be used as an alternative to fibronectin for coating as described in the notes section. In our experience we obtained best results with fibronectin coating.

2.4.3 Collagenase Solution

Dissolve 100 mg of collagenase type I-A (cell culture compatible) in 100 mL of sterile 1X PBS (pH 7.4). Sterilize the collagenase solution by filtering it through 0.22 μm filter. Make 10 mL aliquots of filter-sterilized collagenase solution and store at -20°C. Collagenase solution can be stored at -20°C for up to 2–3 weeks without repeated freeze thawing. Collagenase preparation should be performed in dark and cover the aliquots with aluminum foil till further use. It is however preferable to freshly prepare collagenase solution in dark on the day of isolation of cells.

2.4.4 Endothelial Cell Culture Media

HUVECs can be grown successfully in EGM™ BulletKit™ medium (Lonza, CC3124). It contains EBM basal media and EGM SingleQuots supplement pack. To prepare endothelial cell growth media, add 50 mL FBS to 450 mL EBM basal media. Add

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all growth factors from EGM SingleQuots supplement pack. Also add penicillin (final concentration: 100 U/mL) and streptomycin (final concentration: 100 μg/mL). Gently mix all the media components and filter sterilize the media through 0.22 μm membrane filter (final FBS concentration: 10%). Alternatively, other commercially available media such as Endothelial Cell Medium (ScienCell Research Laboratories, 1001) or Endothelial Cell Growth medium MV2 (Promo Cell, C22121) can also be used for HUVEC culture. [Precaution: It is necessary to heat FBS solution at 56°C for 30 min in a water bath prior to use. One can aliquot serum and store them in -20 °C to avoid multiple freeze thawing. Heating of serum is necessary to inactivate the complement system and to prevent its cytolytic activity.] 2.4.5 Paraformaldehyde for Cell Fixation

Dissolve 4 g of PFA in 50 mL of 1X PBS with constant stirring on a hot plate set at 60°C without boiling the solution. Add few drops of 1 N NaOH until the solution is clear and make up the volume to 100 mL with 1XPBS. 4% PFA can be stored at 4°C for a week. Alternatively, it can be frozen in aliquots and stored at -20°C.

2.4.6 Blocking Solution for Immunofluorescence

Freshly prepare blocking solution (5% FBS) before starting the protocol. Add 50 μL of FBS to 950 μL of 1XPBS and store it at room temperature.

3

Methods

3.1 Umbilical Cord Collection and Processing

1. Researchers should follow applicable ethical and regulatory guidelines for use of human tissues and cells. They should secure relevant Institutional Ethics Committee/Institutional Review Board permissions prior to start of isolation and culture of HUVECs. As a biosafety precaution, the researcher handling the umbilical cord should be immunized against hepatitis B virus. 2. Sterile sample collection boxes (filled with 100 mL of freshly made sample collection buffer) should be placed at 4°C in maternity hospital. It is preferred to have multiple sample collection boxes available in the maternity unit, and each cord should be collected in a separate box. 3. Following successful delivery, umbilical cords (approximately 20–30 cm in length) should be collected and placed in cold sample collection buffer. Ensure that the cords are collected from HIV- and HBV- subjects to minimize risks to the researchers. It is advisable to collect the cords immediately within half an hour of delivery, and isolation of cells should be done within 6 h of cord collection. Transport the collected umbilical cord/s while maintaining 4°C to the experimental lab for further processing.

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4. Before starting the experiment, incubate the endothelial cell culture media and collagenase at 37°C in a water bath for 10–15 min. Frozen collagenase aliquot (-20°C) should be thawed completely and prewarmed at 37°C prior to use. [Precaution: Longer incubation of collagenase at 37°C can reduce its enzymatic activity.] 5. Processing of the cord and other experimental steps should be performed under sterile conditions in the laminar flow hood. Before starting the experiment, wipe the surface of hood with 70% ethanol and switch on the UV light for 15 min. 6. Take the umbilical cord out from the collection box and place it in a beaker filled with sterile 1X PBS. Wash the cord with 1X PBS thrice to clean off the blood from the surface of the cord. 7. Place the cord on the surface of a clean sterile aluminum foil and wipe it dry with the help of tissue/filter paper. Trim/cut off 1 cm of cord from both the ends. This will clearly expose the two umbilical arteries and one umbilical vein. Umbilical vein is the largest vessel in the umbilical cord having larger lumen and thin walls, and it can be easily identified as the vessel from which the dark venous blood oozes out upon trimming of the cord ends. The two umbilical arteries have a smaller lumen with thicker walls. Figure 1b provides schematic representation of the cross section of an umbilical cord to identify the umbilical vein. [Precaution: Aluminum foil and tissue paper should be sterile. Before starting the umbilical cord cell isolation, aluminum foil and tissue paper can be UV sterilized in laminar flow hood. Surface of aluminum foil can also be wiped with 70% ethanol before placing the umbilical cord on it.] 8. After identifying the umbilical vein, insert a 21-gauge butterfly needle into the vein carefully without rupturing and clamp it using a hydrostat scissors on the top. [Precaution: Insert the 21-gauge butterfly needle along with the safety casing so the needle does not pierce the lumen of the vein. It prevents collagenase from entering the subendothelial space. If collagenase enters into the subendothelial space, it will reduce the final yield of endothelial cells.] 9. Take 10 mL of 1X PBS in a syringe and perfuse the umbilical vein with PBS to remove blood and blood clots. Repeat this step until clear PBS starts coming out from the other end of the cord. It usually takes two or three repetitions for complete cleaning of the cord. At the same time, also check if umbilical cord is leaking somewhere along the length. Make sure that the umbilical cord is intact and does not have perforations. If PBS is leaking from these perforations, then discard the cord as it cannot be used for effective isolation of HUVECs.

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10. The maternity staff collecting the umbilical cord can be instructed to collect the umbilical cord blood (as this is done in some hospitals) from one end of the cord instead of punching it at multiple locations along the length of the cord. It is also advisable not to isolate HUVECs from pre-eclampsia or gestational diabetic mothers for regular experiments, since the cells isolated from them exhibit inflammatory phenotype [18–20]. 3.2 Isolation and Culture of HUVECs

1. Place the cord on sterile aluminum foil and wipe it dry with tissue paper. 2. Now with the help of butterfly needle inserted into the umbilical vein, perfuse the umbilical vein with 4 mL collagenase solution, and once it starts coming out from the other end, stop perfusion of the collagenase solution, and clamp the other end of umbilical cord also either with hydrostat clamp or cord clamp as shown in Fig. 1c. 3. Now, fill the umbilical vein with additional collagenase enzyme (usually 10–12 mL of collagenase solution is sufficient for a 20-cm-long umbilical cord). Filling with collagenase will cause distension of the umbilical cord. Gently massage the umbilical cord with fingers throughout the entire length of the cord. [Precaution: Do not remove the syringe attached to butterfly needle after collagenase insertion. The sterile plunger attached to the butterfly needle acts a stopper on its own to prevent backflow of collagenase from the cord. If one tries to remove the syringe, the initial pressure built up inside the cord will stream out collagenase that will lead to less enzyme inside the cord.] 4. Cover the umbilical cord with sterile aluminum foil and place it in an incubator at 37°C. Incubate the cord for 15–20 min. After incubation take out the umbilical cord and massage it again very gently with fingers to detach the endothelial cells from the vessel wall. 5. Take 10 mL of endothelial cell growth media in a sterile disposable clinical grade syringe, and fix it in to the angiocath of butterfly clamp (Fig. 1c). Remove the cord clamp from the distal end of the cord, and collect the collagenase suspension containing endothelial cells in a tissue culture-grade sterile 50 mL falcon tube. Now, gently plunge in endothelial cell growth media into the umbilical vein, and collect the eluate from the distal end into the same falcon tube. 6. Centrifuge the collected cell suspension at 1500 RPM (320 × g) for 5 min in a swinging bucket rotor at room temperature. Following centrifugation, isolated endothelial cells get collected at the bottom of the falcon tube. Discard the supernatant carefully without disturbing the cell pellet.

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7. Gently resuspend the cells in 5 mL of prewarmed endothelial cell growth medium, and seed the cells into two fibronectincoated T-25 flasks at a preferred cell density of 0.7 × 106 cells in each T-25 flask. Incubate the T-25 flasks in cell culture incubator at 37°C temperature, 5% CO2, and 95% humidified air. The average yield from a 20-cm-long umbilical cord is usually 1.5 × 106 cells. 8. The following day, aspirate the media by using sterile pipette and gently rinse off the non-adhered cells and blood cells (if any) with prewarmed sterile 1X PBS. Replenish each of the T-25 flasks with 4 mL of endothelial cell growth media, and place it again in incubator for further incubation. 9. Henceforth, keep changing the cell growth media every 2–3 days, and observe them once a day under a phase contrast microscope till the cells reach confluency. It usually takes 5–7 days for isolated cells to become confluent (as seen in Fig. 2a. This is referred to as passage “0” of HUVECs. Thus, by the end of the seventh day, confluent monolayer of HUVECs is obtained in two T- 25 flasks (2.5 × 106 cells per flask) from a 20-cm-long umbilical cord. 3.3 Trypsinization and Subculture of HUVECs

1. HUVECs (P0) can be subcultured after a confluent monolayer of cells is formed. For subculture of HUVECs, first aspirate the media and wash the cells twice with sterile 1X PBS. 2. Add 0.5 mL of trypsin-EDTA solution to each T-25 flask and shake it gently so that trypsin can cover the full surface of the T-25 flask. Incubate it at 37°C for 3–5 min. Examine the cells under a microscope to ensure that all cells have been detached from the surface of the flask. 3. Add 5 mL of endothelial cell growth media (containing serum) to each flask. Addition of serum containing growth medium will inactivate the trypsin. Gently aspirate with the help of a sterile pipette the cell suspension five to six times to dissociate the cell aggregates. Now transfer the homogenous cell suspension into a 15 mL falcon tube, and centrifuge them in a swinging bucket rotor at 1500 RPM (294 × g) for 5 min at room temperature. Cells would be collected at the bottom of the tube following centrifugation. 4. Discard the supernatant and resuspend cells in 1 mL of endothelial cell growth medium. Count the cells using a hemocytometer and seed approximately 0.7 × 106 cells as initial seeding density into each fibronectin coated T-25 flask containing 4 mL of endothelial cell growth media and place them in cell culture incubator. In general, cells from one confluent T-25 flask can be split and seeded into three fresh fibronectin-coated

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Day 2

Day 1

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Day 7

B Negative control

VE-cadherin

20µm

DAPI

eNOS

20µm

20µm

vWF

Merge

10µm

Dil-LDL uptake

50µm

C

PBMCs

200µm

Ulex lectin

50µm

HUVECs (batch-1) HUVECs (batch-2)

200µm

200µm

Fig. 2 (a) Phase contrast image of cultured HUVECs at different days of culture. (b) Validation of endothelial phenotype of cultured HUVECs through immunofluorescence imaging of endothelial markers: VE-cadherin, eNOS, vWF, Lectin staining, and dil-acetylated LDL uptake. Images for vWF were acquired through a confocal microscope. (c) Capillary tube formation by primary HUVECs on Matrigel which is not seen for the freshly isolated peripheral blood mononuclear cells (PBMCs). Thus, PBMCs were used as a negative control

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T-25 flasks. Every alternate day, replenish the media and check for confluency under phase contrast microscope. It usually takes 4–5 days for cells to attain confluency. This cell monolayer will constitute passage 1 (P1) of HUVECs. 5. If the researcher wishes to perform experiments with P1 of HUVECs, then HUVECs (P0) can be directly seeded as per experimental requirements after trypsin treatment into the desired culture dishes. A good seeding density is when approximately 40% of the surface area of the culture dish is covered at the time of seeding itself. HUVECs do not grow very well at very low cell density. 6. HUVECs can be conveniently subcultured after isolation for up to 6 weeks. It is advisable to perform experiments with second to fifth passage of culture. After this, one may notice phenotypic changes in cultured endothelial cells. 3.4 Characterization and Validation of HUVECs

While standardizing the protocol for isolation and culture of HUVECs, researchers can resort to morphological as well as functional validation of endothelial cells to confirm their phenotype and purity of culture.

3.4.1 Morphological Validation

A confluent monolayer of endothelial cells will consist of a homogenous population of polygonal cells with centrally placed nucleus. These cells when visualized through phase contrast microscope will exhibit a cobblestone appearance at confluence as seen in Fig. 2a.

3.4.2 Immunofluorescence Staining for Endothelial Markers

Endothelial cells can be characterized by determining the expression of endothelial-specific markers such as VE-cadherin, von-Willebrand factor (vWF), and eNOS [21, 22]. Other markers that can be used for confirmation of endothelial phenotype are E-selectin, CD31, and VEGF receptors 1 and 2 [23]. 1. Culture HUVECs on fibronectin-coated glass bottom chamber slides which are compatible for imaging. Once a tight HUVEC monolayer is formed, aspirate the media, and wash the cells with 1X PBS. For staining of cells with VE-cadherin, it is necessary that the cells reach confluence, as it is best seen at intercellular junctions. 2. Fix the cells with 4% PFA for 20 min at room temperature. After fixation, wash the cells with 1X PBS two times. 3. Permeabilize the cells with 0.25% Triton X-100 (molecular biology grade) in 1X PBS for 10 min on ice. Wash the cells with 1X PBS two times. 4. Add blocking solution (5% FBS in 1X PBS) and incubate at 37 °C for 1 h on a rocking platform.

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5. Following blocking, add primary antibody at specified dilution and incubate at 4°C overnight on a rocking platform. Dilute the antibody in blocking solution as listed below for different antibodies. 6. Next day, wash the cells with 1X PBS three times. Add fluorescence conjugated secondary antibody and incubate in dark for 1 h at room temperature as per the details listed below. Dilute the secondary antibody in the blocking solution. 7. Aspirate the secondary antibody and wash with 1X PBS three times followed by counterstaining with DAPI (1 μg/mL) and take the images in fluorescence microscope (Fig. 2b). 8. Primary antibody dilutions: • eNOS (dilution 1:500). • VE-cadherin (dilution 1:500). • vWF (dilution 1:250). Secondary antibody dilutions: • Alexa 594 secondary anti-mouse (dilution 1:10000). • Alexa 488 secondary anti-rabbit (dilution 1:10000). 3.4.3 Dil-Acetylated LDL Uptake Assay and Lectin Staining

1. Scavenger receptor SERC present on endothelial cells assists in LDL uptake [24, 25]. Thus, staining of HUVECs with fluorescently tagged LDL and its internalization allows for functional validation of endothelial cells. For Dil-Ac-LDL uptake assay, culture HUVECs either in glass bottom chamber slides or in microscope compatible 35 mm cell culture-grade dishes. Incubate the confluent culture of HUVECs with 1–10 μg/mL Dil-Ac-LDL dye in endothelial cell growth media for 4 h at 37°Cin cell culture incubator. Wash the cells with 1X× PBS and capture the images using green excitation filter of an immunofluorescence microscope (Fig. 2b). 2. Endothelial cells in human blood vessels express blood group ABH antigens as per the donor blood type [26, 27]. However, endothelial cells from umbilical vein predominantly express the H-antigen, while the A and B antigens are expressed only in the regions proximal to the fetus [28]. Lectin, a Ulex europaeusderived agglutinin, is a glycoprotein which binds to alpha-L fucosyl residues of oligosaccharides of the H-antigen present on endothelial cell membrane. For lectin staining, incubate the confluent culture of HUVECs with 10 μg/mL Ulex lectin (FITC conjugated) in endothelial basal media for 30 min at 37°C in cell culture incubator. Wash with 1X PBS gently and capture the images using blue excitation filter of an immunofluorescence microscope (Fig. 2b). Since these antigens are not present in fibroblasts and smooth muscle cells, they stain weakly with lectin.

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3.4.4 Tube Formation Assay

Endothelial cells exhibit angiogenic potential by undergoing proliferation, migration, and tubulogenesis [29, 30]. In a 3D solubilized extracellular matrix, cultured endothelial cells form a network of capillary tubes. HUVECs can also be characterized for their angiogenic potential by performing in vitro tube formation assay. A brief protocol for the same is mentioned below: 1. Thaw Matrigel (Corning® Matrigel; growth factor reduced, Phenol Red-free, Cat No: 356230) matrix as recommended by the manufacturer. Mix it to homogeneity using prechilled sterile pipettes and tips. 2. Place 96-well culture plates on ice and add 50 μL of chilled Matrigel matrix (10 mg/mL) per well. This quantity is sufficient to cover the entire growth surface of the well. 3. Avoid air bubbles in Matrigel matrix while pipetting the liquid into each well. 4. Trypsinize the HUVEC monolayer and resuspend the cells in endothelial growth medium. 5. Add 15,000 cells/well and incubate the plate for 16 h at 37°C, 5% CO2 cell culture incubator. One can also add growth factors such as VEGF (10 ng/mL) to enhance tube formation ability of cultured HUVECs. 6. Take images at 4X using inverted phase contrast microscope (Fig. 2c). [Precaution: Matrigel polymerizes very fast at temperature 22–35 °C. Thaw Matrigel overnight at 4 °C on ice and use precooled pipette tips, plates, and tubes during Matrigel preparation and coating. It is advisable to make the aliquots of Matrigel and stored to avoid multiple freeze thaw.]

3.5

4

Troubleshooting

Various difficulties that one may encounter during isolation and culture of HUVECs and the possible reasons for them are listed in Table 1.

Notes 1. A written pre-informed consent must be obtained before collection of umbilical cords. 2. Sample collection buffer should be sterile. Sterilize the sample collection boxes by autoclaving every time before use. Collect only one umbilical cord in one sample collection box. Do not collect more than one cord in same box. Since endothelial cells express HLA-DR antigens, pooling of cords may not be possible all the time. Collecting more than one cord in same box will also increase the risk of contamination.

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Table 1 Various difficulties that one may encounter during isolation and culture of HUVECs and the possible reasons for them S. No. Observations

Likely reasons

1.

Few or no cells in the eluate after collagenase treatment

1. This could be due to an old preparation of enzyme 2. This could be due to poor quality of collagenase enzyme

2.

Poor adherence of cells on coated dishes after 24 hours

1. Poor quality of ECM protein 2. Inappropriate functioning of CO2 incubator or inappropriate humidity conditions 3. Cells isolated from patient samples such as pre-eclampsia and gestational diabetes 4. Absence of Ca2+ ions in culture media 5. If cords from different subjects were pooled, then it could be due to HLA-DR incompatibility

3.

Cells do not reach confluency within 10 days

1. Poor quality of culture media such as inappropriate pH 2. Contamination of culture with mycoplasmas, bacteria, or fungi 3. HLA incompatibility between pooled cords

4.

Many detached cells

1. Unhealthy cords 2. Isolation of cells done after 6 h of delivery 3. Contamination

5.

Elongated slender spindle-shaped cells

1. Contamination of cell preparation with fibroblasts or smooth muscle cells

6.

Filamentous hair-like strands in culture dish

This indicates fungal contamination

7.

Cloudy appearance of culture medium

This indicates bacterial contamination

8.

Branched ovoid particles budding off

This indicates yeast contamination

3. Avoid collecting umbilical cords from women with HBV+ or HIV+ infection. 4. Isolation of cells from the umbilical cords should be done within 6 h of delivery. Yield of isolated HUVECs will decrease if the umbilical cord has been stored for longer time. Longer storage time of umbilical cord also enhances the risk of contamination. 5. At the time of isolation, if cord is leaky or damaged, then it is better to discard them. 6. It is important that solutions used for preparation of media, collagenase, fibronectin solution, etc. should be endotoxinfree. This is because endotoxin can interfere with endothelial proliferation and can enhance their inflammatory phenotype.

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7. Do not confuse the umbilical vein with the two arteries. Umbilical vein is the widest among all and has thin vessel wall. 8. Umbilical cord should be cleaned properly by PBS. Flush the cord with PBS until all red blood cells are removed. Sometimes, umbilical cord is clogged with multiple blood clots and does not allow PBS to pass through. At the time of cord collection, gently squeeze the blood out of the umbilical cord. This will avoid the clogging of the cord with blood clots. 9. For collagen coating, add 2.5 mL of rat tail collagen solution prepared in 0.02 M acetic acid at a concentration of 50 μg/mL, to each T-25 flask and incubate at 37°C for 2 h in cell culture incubator. After 2 h, aspirate the excess collagen solution using sterile pipette and rinse the coated surface with 1X PBS twice. Collagen-coated flask can be stored at 4°C for a week. For gelatin coating, add 2 mL of 0.2% gelatin solution to each T-25 flask and allow it to air dry for 1 h at room temperature in cell culture hood. After 1 h, aspirate the excessive gelatin solution by using sterile pipette tip. Do not over dry the gelatin coating as it disrupts the gelatin structure. 10. It is always better to use freshly made collagenase. Alternatively, collagenase can be aliquoted and stored at -20°C. Do not freeze thaw multiple times. 11. Do not incubate for more than 20 min with collagenase solution. Longer incubation leads to contamination with fibroblasts and smooth muscle cells. Likewise, multiple clamping or rigorous massaging of the cord will disrupt the basement membrane, thereby allowing dissociation of underlying smooth muscle cells. 12. Do not allow cells to be overconfluent, as it will lead to cell death and detachment. 13. As mentioned, cultured fibroblasts and smooth muscle cells appear as long slender spindle-shaped cells that grow in multiple parallel layers. 14. HUVECs can survive in serum-free medium without losing their phenotypic property for up to 12 h. 15. Microbial contamination is a huge challenge during HUVEC isolation. Bacterial contamination can be avoided by using penicillin-streptomycin and gentamicin in culture media and endothelial growth medium. To mitigate fungal contamination, ketoconazole (10 μg/mL) can be used in collection buffer. Maintain complete sterile environment at the time of HUVEC isolation and culture. HUVECs must be checked for mycoplasma contamination also. This can be detected through Hoechst 33258 staining. If any contamination is present in cultured HUVECs, it should be immediately discarded.

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16. While handling human tissue, always use gloves, lab coat, mask, and head cap for personal safety. 17. After umbilical cord processing, tissue must be discarded as per biosafety guidelines. 18. Primary cells that proliferate but fail to become confluent within 7 days can be pushed to grow better by detaching them and reseeding them in fresh fibronectin-coated flasks. 19. HUVECs can be successfully cultured and passaged up to 6 weeks. However, expression of multiple endothelial markers decreases upon subculturing of HUVECs after six passages [31]. 20. Endothelial growth media available from commercial suppliers such as PromoCell or Lonza are MCDB131 based, and they contain growth factors such as VEGF which may not be preferred by some researchers. One can also use endothelial growth medium as reported by others [32]. To cut down long-term costs, one can also grow endothelial cells in M199 medium supplemented with 20% fetal calf serum (FCS) and antibiotics such as penicillin and streptomycin, but in our experience, best results are obtained when the medium is supplemented with endothelial cell growth supplements as seen for the media available from Lonza or PromoCell.

Acknowledgments The authors acknowledge funding received from the Department of Biotechnology (DBT) [File# BT/PR12547/MED/30/1456/ 2014] and Science and Engineering Research Board (SERB) [File# EMR/2015/000704], Government of India for their work. References 1. Jaffe EA (1987) Cell biology of endothelial cells. Hum Pathol 18(3):234–239 2. Michiels C (2003) Endothelial cell functions. J Cell Physiol 196(3):430–443 3. Pober JS, Sessa WC (2007) Evolving functions of endothelial cells in inflammation. Nat Rev Immunol 7(10):803–815 4. Sandoo A, van Zanten JJ, Metsios GS, Carroll D, Kitas GD (2010) The endothelium and its role in regulating vascular tone. Open Cardiovasc Med J 4:302–312 5. Davignon J, Ganz P (2004) Role of endothelial dysfunction in atherosclerosis. Circulation 109(23 Suppl 1):III27–III32 6. Rajendran P, Rengarajan T, Thangavel J, Nishigaki Y, Sakthisekaran D, Sethi G,

Nishigaki I (2013) The vascular endothelium and human diseases. Int J Biol Sci 9(10): 1057–1069 7. Liao JK (2013) Linking endothelial dysfunction with endothelial cell activation. J Clin Invest 123(2):540–541 8. Michiels C, Arnould T, Remacle J (2000) Endothelial cell responses to hypoxia: initiation of a cascade of cellular interactions. Biochim Biophys Acta 1497(1):1–10 9. Schwartz SM (1978) Selection and characterization of bovine aortic endothelial cells. In Vitro 14(12):966–980 10. Fehrenbach ML, Cao G, Williams JT, Finklestein JM, Delisser HM (2009) Isolation of

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murine lung endothelial cells. Am J Physiol Lung Cell Mol Physiol 296(6):L1096–L1103 11. Ruck T, Bittner S, Epping L, Herrmann AM, Meuth SG (2014) Isolation of primary murine brain microvascular endothelial cells. J Vis Exp no. 93:e52204 12. Jaffe EA, Nachman RL, Becker CG, Minick CR (1973) Culture of human endothelial cells derived from umbilical veins. Identification by morphologic and immunologic criteria. J Clin Invest 52(11):2745–2756 13. Baudin B, Bruneel A, Bosselut N, Vaubourdolle M (2007) A protocol for isolation and culture of human umbilical vein endothelial cells. Nat Protoc 2(3):481–485 14. Cheung AL (2007) Isolation and culture of human umbilical vein endothelial cells (HUVEC). Curr Protoc Microbiol Appendix 4:Appendix 4B 15. Onat D, Brillon D, Colombo PC, Schmidt AM (2011) Human vascular endothelial cells: a model system for studying vascular inflammation in diabetes and atherosclerosis. Curr Diab Rep 11(3):193–202 16. Bachetti T, Morbidelli L (2000) Endothelial cells in culture: a model for studying vascular functions. Pharmacol Res 42(1):9–19 17. Bishop ET, Bell GT, Bloor S, Broom IJ, Hendry NF, Wheatley DN (1999) An in vitro model of angiogenesis: basic features. Angiogenesis 3(4):335–344 18. Giri H, Chandel S, Dwarakanath LS, Sreekumar S, Dixit M (2013) Increased endothelial inflammation, sTie-2 and arginase activity in umbilical cords obtained from gestational diabetic mothers. PLoS One 8(12):e84546 19. Pantham P, Aye IL, Powell TL (2015) Inflammation in maternal obesity and gestational diabetes mellitus. Placenta 36(7):709–715 20. Redman CW, Sacks GP, Sargent IL (1999) Preeclampsia: an excessive maternal inflammatory response to pregnancy. Am J Obstet Gynecol 180(2 Pt 1):499–506 21. Jaffe EA, Hoyer LW, Nachman RL (1974) Synthesis of von Willebrand factor by cultured human endothelial cells. Proc Natl Acad Sci USA 71(5):1906–1909

22. Larrivee B, Karsan A (2005) Isolation and culture of primary endothelial cells. Methods Mol Biol 290:315–329 23. Garlanda C, Dejana E (1997) Heterogeneity of endothelial cells. Specific markers. Arterioscler Thromb Vasc Biol 17(7):1193–1202 24. Voyta JC, Via DP, Butterfield CE, Zetter BR (1984) Identification and isolation of endothelial cells based on their increased uptake of acetylated-low density lipoprotein. J Cell Biol 99(6):2034–2040 25. Adachi H, Tsujimoto M, Arai H, Inoue K (1997) Expression cloning of a novel scavenger receptor from human endothelial cells. J Biol Chem 272(50):31217–31220 26. Szulman AE (1962) The histological distribution of the blood group substances in man as disclosed by immunofluorescence: II. The H antigen and its relation to A and B antigens. J Exp Med 115(5):977–996 27. Szulman AE (1964) The histological distribution of the bloop group substances in man as disclosed by immunofluorescence: III. The A, B, and H antigens in embryos and fetuses from 18 mm in length. J Exp Med 119:503–516 28. Szulman AE (1972) The A, B and H bloodgroup antigens in human placenta. N Engl J Med 286(19):1028–1031 29. Cantelmo AR, Brajic A, Carmeliet P (2015) Endothelial metabolism driving angiogenesis: emerging concepts and principles. Cancer J 21(4):244–249 30. Lamalice L, Le BF, Huot J (2007) Endothelial cell migration during angiogenesis. Circ Res 100(6):782–794 31. Goldsmith JC, McCormick JJ, Yen A (1984) Endothelial cell cycle kinetics. Changes in culture and correlation with endothelial properties. Lab Investig 51(6):643–647 32. Sow RC (2012) Introduction to Cell culture: Culture of human endothelial cells from umbilical veins. Methods Mol Biol 806: 265-274.

Chapter 13 Microvascular Endothelial Glycocalyx Surface Layer Visualization and Quantification Natascha G. Alves and Jerome W. Breslin Abstract As a primary interface between the blood and underlying vascular wall, the endothelial glycocalyx layer is common to all blood vessels and covers the luminal surface of all endothelial cells. The endothelial glycocalyx has important roles as a regulator of microvascular endothelial functions such as mechanotransduction, leukocyte adhesion, and microvascular permeability. Disruption of the molecular structure of the endothelial glycocalyx disturbs physiological, and hemodynamic processes associated with the microvascular wall leads to microvascular hyperpermeability. Studying the glycocalyx is challenging because cultured cells present aberrant glycocalyx structure and tissue fixation techniques lead to the degradation and loss of this fine and delicate layer. Therefore, studying the glycocalyx requires in vivo imaging of the microcirculation. Here we describe two techniques for direct imaging and assessment of the glycocalyx surface layer integrity using intravital microscopy (IVM), a method widely used in the study of the dynamic changes that occur in the microcirculation during inflammation or injury. Key words Glycocalyx, Endothelial surface layer, Dye exclusion, BSI-Lectin, Intravital microscopy

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Introduction The luminal surface of the endothelium is coated with a thin gel-like layer composed of extracellular proteoglycans and glycoproteins associated with glycosaminoglycans. This endothelial glycocalyx layer (EGL) is loosely bound to the luminal (apical) endothelial cell membrane [1, 2]. The EGL composition and thickness are unevenly distributed throughout different vascular beds. For instance, in the mesenteric microcirculation, particularly in the highly permeable venules, the EGL varies in thickness and composition, while in the aorta it is more evenly distributed [3–5]. The EGL has been demonstrated to regulate a variety of endothelial functions. These include microvascular permeability, nitric oxide bioavailability, immune cell-endothelium interactions, and shear stress mechanotransduction [6–8]. Degradation of the glycocalyx is associated with ischemia-reperfusion injury [9, 10] and with

Binu Tharakan (ed.), Vascular Hyperpermeability: Methods and Protocols, Methods in Molecular Biology, vol. 2711, https://doi.org/10.1007/978-1-0716-3429-5_13, © Springer Science+Business Media, LLC, part of Springer Nature 2024

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increased microvascular permeability in in vivo models of traumatic injury such as hemorrhagic shock, as well as in severely injured trauma patients [5, 11]. Intravital microscopy (IVM) enables real-time visualization of the microvasculature and measurements of several biological processes in the microcirculation, including changes in vessel diameter, blood flow, leukocyte rolling/adhesion, and microvascular permeability [12]. The recognition of the glycocalyx as an important component of the microcirculation and regulator of vascular endothelial function has also been determined using specific IVM methods [13, 14]. Direct study of the glycocalyx has been limited by its small dimensions and by standard tissue processing techniques [7, 9]. For instance, the EGL is invisible when using transmitted light microscopy, and tissue fixation and processing generally result in degradation of the glycocalyx [7]. However, specific IVM techniques have been developed to quantitate the thickness and integrity of the EGL in live tissues. Both of these glycocalyx visualization techniques require fluorescence microscopy [5]. Both of these methods are complimentary and relatively direct measurements of glycocalyx integrity. Other, less direct methods are based on the ability of the EGL to exclude circulating elements from the endothelial surface. For instance, the ability of circulating fluorescent microspheres to access and interact with the endothelium has been measured as an indirect indicator of EGL integrity [7]. The methods to assess glycocalyx integrity may be combined with disease or injury models for real-time assessment of the dynamic changes that happen to the glycocalyx during pathophysiological conditions where microvascular hyperpermeability is present. The rat mesentery is a good option for permeability studies because it is quickly affected by insults that induce hyperpermeability such as ischemia-reperfusion injury and acute inflammation [15]. The mesentery is a thin tissue that allows transillumination, making it ideal for studying the exclusion properties of the glycocalyx when techniques involving fluorescent-labeled tracer molecules are applied [16]. Other vascular beds such as the cremaster muscle are also well-suited for the study of the glycocalyx in microvascular hyperpermeability because they are relatively translucent, can be accessed with relative ease, and generally do not have many large movements. This being said, there are also techniques for IVM imaging of the free-moving rodent lung that can be adapted for glycocalyx studies [7, 17]. It is important to note that the uptake and binding of lectin to the endothelial glycan layer depends on blood flow and is convection limited [18]. Therefore, lectin uptake over time differs among vessel types and should be accounted for when imaging venules, arterioles, or capillaries. Besides that, models that affect blood flow may influence lectin uptake. When using such models, ensure to select areas or frames that have similar and steady flow rate for

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imaging. It is also important to consider that the FITC-lectin method does not allow measurement of changes in the composition of the glycocalyx. To this end, it is recommended to quantify the presence of glycocalyx components like syndecan-1 and heparan sulfate – two of the main components of the glycocalyx – in the plasma by methods such as ELISA, as an indicator of composition changes [18, 19]. To date, BSI-lectin has not been shown to have any direct effect on microvascular function. However, it has been observed that when lectins are conjugated with other ligands, such as ferritin, modification of the glycocalyx layer leads to altered microvascular function in the mesentery [20].. The dye-exclusion method is an additional technique that can be used to quantify thickness of the glycocalyx surface layer on the endothelium. This technique arose from the observation of an “exclusion zone” on the microvascular wall surface, such that the measurement of luminal diameter of microvessels using an intravenous fluorophore-bound tracer with a 70 kDa molecular weight or greater yields a narrower value than when measuring the distance between vessel walls by brightfield microscopy in the exact same area [21, 22]. The composition and distribution of the glycocalyx layer varies depending on the vascular bed and vessel type. For instance, the EGL is unevenly distributed on postcapillary venules of the mesentery, whereas it is evenly distributed on the aorta [3, 4]. Therefore, it is important to note that the dye-exclusion technique only provides a rough estimation of the thickness of the EGL on vascular beds where it has uneven distribution, like the mesentery. Because of that, at least two imaging techniques should be employed to achieve an accurate estimation of EGL integrity. In this chapter, we describe two procedures in which IVM using epifluorescence illumination is used to visualize and quantitate the endothelial glycocalyx integrity and thickness in vivo. The two methods include (1) quantification of the intensity of Bandeiraea simplicifolia (BSI)-lectin conjugated to fluorescein isothiocyanate (FITC) within microvessels, and (2) the Dye-Exclusion assay. We describe these methods in the microcirculation of the rat mesentery, which is well suited for intravital microscopic observation of microvessels.

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2.1 Vascular Cannulation and IVM Preparation

1. Vascular catheter either made from polyethylene 50 tubing [23] or purchased from a commercial source (SAI Infusion Technologies). 2. Fine scissors, available from Fine Science Tools (FST), catalog no. 14958-11. 3. Tissue forceps (FST 11021-14).

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4. Microdissection forceps (Roboz RS-5069).. 5. Fine forceps (Roboz RS-5137 T). 6. Vannas scissors (FST 15000-00). 7. Ultra-fine hemostats (FST 13021-12). 8. 1-cc syringes. 9. 23-gauge needles. 10. Betadine scrub. 11. Chlorhexidine gluconate scrub. 12. Alcohol. 13. 0.9% sodium chloride USP. 14. 10-cc syringes. 15. Gauze sponges. 16. Cotton-tipped applicators. 17. Ethicon FS-2 suture and needle (Johnson & Johnson). (a) Alternative: stainless steel wound clips (Roboz RS-9258). 18. Ethicon 4–0 silk, black, braided suture without needle (Johnson & Johnson). 19. Isoflurane USP and vaporizer. 20. Cotton-tipped applicators. 21. Akwa tears ophthalmic ointment (Akorn). 22. Fur trimmers. 23. 2 × 2 gauze sponges. 24. Lactated ringers (LR) USP. 25. Heating pad. 26. Rat stage from IVM. 27. Peristaltic pump. 28. Heating coil (Radnoti). 29. Water bath circulator. 2.2 Glycocalyx Visualization with BSILectin and DyeExclusion Assay

1. Upright fluorescent microscope (e.g., Nikon E600FN). 2. Euthasol – pentobarbital sodium and phenytoin sodium. 3. Bandeiraea simplicifolia (BSI)-lectin conjugated to fluorescein isothiocyanate (FITC) (FITC-lectin; Millipore Sigma). 4. FITC-150 kDa dextran (FITC-Dx150) (Millipore Sigma). 5. 0.9% sodium chloride USP. 6. 1-cc syringe.

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Methods

3.1 Blood Vessel Cannulation

Prior to the beginning of the IVM and glycocalyx integrity assessment procedures, an intravenous catheter must be placed in order to administer the fluorophore-conjugated indicators that will be used in the protocol. Typically, a polyethylene catheter is implanted into one of the external jugular veins, although other veins may also be used. If monitoring of blood pressure during the experiment is also desired, then placement of another catheter into the contralateral common carotid artery can also be performed as outlined previously [23]. Animals should be anesthetized throughout the entire protocol. Anesthesia can be achieved with isoflurane or other anesthetics such as ketamine/xylazine. Here, we describe the use of isoflurane: 1. Prepare the venous catheter by flushing it and filling it with sterile saline solution using a 1-cc syringe and 23-gauge needle. Ensure there are no air bubbles within the catheter (see Note 1). 2. To anesthetize with isoflurane, place the rat in an induction chamber and open the isoflurane vaporizer to 3–4%. Open the USP-grade oxygen cylinder and set the flow rate to 1 liter (L) per minute (min). 3. Ensure the rat is anesthetized by lightly pinching its toes with forceps before removing it from the chamber. 4. Reduce the isoflurane to 2.0–2.5% (see Note 2) and redirect to the nosepiece. 5. Place the rat in a dorsally recumbent position on a warm heating pad. Insert the nose into the nosepiece to maintain isoflurane delivery. 6. Using animal fur trimmers, shave about a 2 × 2 cm area of fur from the ventral neck area. Increase or decrease the shaved area depending on the size of the rat. 7. Using gauze or cotton-tipped applicators, clean the shaved area by applying 70% ethanol once, chlorohexidine surgical scrub once, and povidone-iodine scrub once. 8. Make a small incision (about 1 cm) in the ventral neck along the medial long axis. 9. Using ultra-fine hemostats, carefully isolate either the left or right external jugular vein (see Note 3). 10. Using a small silk suture (4-0) without needle, tie the brain side vessel with two simple suture knots. 11. Place a strip of suture (about 8 cm long) under the carotid vessel on the heart side. Do not tie this suture yet.

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Fig. 1 Implantation of venous catheter. (a) After isolating the jugular vein and placing two sutures underneath it, the suture closer to the head is used to tie off the vessel, and then a small nick incision is made on the vessel as shown. (b) The venous catheter is then inserted into the nick and advanced up to the suture retention bubble. (c) Both sutures are then tied around the catheter as shown, using square knots. (d) The loose strands from both of the square knots can then be tied together to further secure the catheter in place

12. Using Vannas micro-scissors, create a small incision in the jugular vein between the suture knot and the untied suture (Fig. 1a). 13. With one hand, open the incision using micro-dissecting forceps. With the other hand, hold the tip of the catheter using extra fine forceps, and feed the catheter into the incision, about 2 cm into the vessel, up to the catheter’s suture retention bubble (Fig. 1b). 14. Anchor the catheter to the vessel by making two square knots using the sutures placed earlier (Fig. 1c). 15. Check for adequate blood flow into the catheter by observing if the catheter (or syringe if catheter is not clear) starts filling up with blood with slight negative pressure. Flush the catheter with a little bit of saline to remove blood from inside it and avoid blood clots. After this check, if needed adjust the catheter slightly until blood enters easily. After confirming good placement, turn the three-way stopcock toward the catheter so it is “off.” 16. Secure the catheter further by tying the two knots on either side of the suture retention bubble on the catheter can also be tied together to secure the catheter in place (Fig. 1d). 17. Close the incision in the skin with simple suture knots or wound clips so that body heat is not lost through the incision during the experiment.

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IVM Preparation

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1. Place the rat on a heating pad (37 °C), and shave the ventral abdominal fur using rodent hair clippers. 2. Using gauze sponges, clean the shaved area by applying 70% ethanol once, chlorohexidine surgical scrub once, and povidone-iodine scrub once. 3. Perform midline laparotomy: make a vertical incision of about 2.5–3.0 cm long in the middle of the shaved area. 4. Insert one 2 × 2 gauze sponge into the left side of the incision and one into the right side. This prevents additional abdominal contents to come into direct contact with the incision. 5. Using cotton-tipped applicators dipped in saline, begin to remove the small intestine and associated mesentery from the abdominal cavity. Do this by holding the intestinal walls (do not touch the mesentery) with the cotton-tipped applicators and carefully pulling the small intestines out (see Note 4). 6. Turn the rat on its side and transfer onto the IVM stage (see Note 5). 7. Using the cotton-tipped applicators, splay the mesentery flat on the stage. Do this by touching the intestinal walls and be careful not to break the microvessels of the mesentery. Apply some warm saline solution to the mesentery windows to prevent them from drying (see Note 6). 8. Transfer the rat on the stage to the microscope. 9. When setting up on the microscope, keep the mesentery windows superfused with warm Ringer’s solution dripping over the intestines using a peristaltic pump at about 1 mL/min. The Ringer’s solution should be pumped through a heating coil heated to 37 °C by a circulating water bath. Remove excess fluid from the stage using a vacuum line. 10. Turn on all components of the intravital microscope system in preparation of performing either the FITC-BSI lectin procedure or the dye-exclusion assay.

3.3

IVM: FITC-Lectin

Direct visualization of the endothelial glycocalyx layer is possible using fluorescently labeled lectins that bind specific sugar moieties of the glycosaminoglycans that compose the EGL [16]. Here we use FITC-conjugated Bandeiraea simplicifolia (BSI) lectin (FITClectin) for direct glycocalyx visualization. The integrity of the glycocalyx is estimated by quantifying the endothelial glycan layer concentration, which is indicated by the amount of endothelial surface-bound lectin. 1. Prepare a bolus dose of FITC-lectin by dissolving FITC-lectin in 1 mL of warm saline solution. Calculate the amount of BSIlectin based on a dose of 6.25 mg/kg.

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2. Select one mesenteric window and observe it under the microscope. Select an area with a few postcapillary venules that have steady blood flow to take images (see Note 7). Ideally, at least three different venules in each experiment should be selected for imaging. A single frame usually displays at least three venules. 3. Using a 1-cc syringe and 23-gauge needle inserted in the loose end of the jugular catheter, begin the administration of the FITC-lectin bolus solution. This should be done slowly over 1–2 min. 4. Allow 10–15 min for systemic distribution and adhesion of the FITC-lectin to the luminal side of vessel walls. 5. Begin imaging of the selected area using an upright fluorescent microscope. Fluorescent images should be taken with a 10× objective at 488 nM excitation. 6. Continue capturing images at consistent intervals, over the desired period of time. 7. Upon completion of imaging, euthanize the animal by administering Euthasol through the jugular catheter (0.1 mL/450 g) or another approved method in accordance with local regulations. 8. Use the fluorescent images to quantify the endothelial surface glycan concentration. 3.4 FITC-Lectin Image Analysis

1. Analyze the fluorescent images using image analysis software such as FIJI/ImageJ (Fig. 2). 2. Manually trace a measurement path along the curvature of the microvessel wall, using the bound FITC-lectin as reference (see Note 8). 3. Generate a radial measurement line normal to the beginning of the path that spans the entire microvessel width and beyond. 4. The Analyze>Plot profile function in FIJI/ImageJ can be used to generate the intensity distribution in each pixel along the measurement line. 5. The average intensity of the pixels along the line can also be obtained by simply using the Measure function and having mean intensity set as a measurement. 6. In images where both vessel walls are clearly in focus, average the intensities of lines drawn on both vessel walls.

3.5 IVM: DyeExclusion Assay

This technique is based on the knowledge that bulky, fluorescently labeled, vascular markers such as dextrans are excluded from the endothelial surface by the glycocalyx [21]. The thickness of the glycocalyx is estimated by measuring the anatomical width of the

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Fig. 2 Measurement of FITC-lectin intensity on microvascular walls. (a) Example image of rat mesenteric microcirculation after intravenous injection of 6.25 mg/kg FITC-lectin. Two regions of interest (ROI) were drawn on a venule (blue) and arteriole (red). Panels b and c show plots of the intensities of the FITC-lectin labeling along the length of the two ROI lines, generated using FIJI/ImageJ. For both ROI lines, the intensities are quite variable along the length of the vessel wall, hence the need to obtain an average along some length of vessel and the need to evaluate more multiple venules or arterioles

microvessel and the width of the fluorescent column created by the tracer, which is excluded from the EGL. Dextrans 70 kDa or 150 kDa are commonly used for this assay. Here we suggest the use of FITC-dextran with an average 150-kDa molecular weight (FITC-Dx150) for the dye-exclusion method to estimate glycocalyx thickness. 1. Prepare a bolus dose of FITC-Dx150 in 1 mL of warm normal saline to achieve a 100 mg/kg dose. 2. Select one mesenteric window and observe it under the microscope. Select an area with postcapillary venules that have steady blood flow to take images (see Note 7). (a) At least three different venules in each experiment should be selected for imaging. A single frame usually displays at least three venules. 3. Using a 1 cc syringe and 23-gauge needle inserted in the loose end of the jugular catheter, administer the bolus of FITCDx150 over 1–2 min. 4. Allow 5–10 min for systemic distribution of the FITC-Dx150 and for the fluorescence to reach a steady level. 5. Begin imaging the selected area: capture one brightfield image and one fluorescent image (at 488 nM excitation). Be careful not to move the rat or the mesenteric window out of frame when capturing images. 6. Continue capturing images at consistent intervals, over the desired period of time.

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Fig. 3 Measurement of endothelial glycocalyx layer thickness with the dye-exclusion method. A brightfield image and a fluorescent image of infused Texas Red-Dextran-70 kDa (TR-Dx70) were obtained of the same venule. The distance between the endothelial walls and of the intravascular tracer was determined using FIJI/ ImageJ as shown in the line intensity plots. The difference between these distances, divided by two, represents the average glycocalyx layer thickness. Because this value can be variable along the length of the vessel, several measurements should be made and averaged for a given vessel segment

7. Upon completion of imaging, euthanize the animal by administering Euthasol through the jugular catheter (0.1 mL/450 g) or another approved method in accordance with local regulations. 8. Use the brightfield and fluorescent images to estimate the thickness of the glycocalyx layer. 3.6 Dye-Exclusion Assay Image Analysis

Estimate the widths of the fluorescent column using the fluorescent images, and of the microvessel inner diameter using the brightfield images, as described below (Fig. 3). 1. Using the brightfield image, measure the width of the microvessel by tracing three perpendicular lines along the longitudinal axis of the selected vessel. Start each line on the outer edge of the dark band near the vessel wall, and stop on the outer edge of the dark band near the opposite vessel wall. 2. Average the length values of those three lines to obtain the mean vascular diameter. 3. Using the fluorescence images, trace three lines that span the entire width of the fluorescent column and beyond.

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4. Generate a sigmoidal curve to display the profile of average intensity of each line, and measure the distance between the beginning and the end of the curve. 5. Average the distances obtained from the three lines to obtain the mean width of the fluorescent column. 6. Subtract the mean value of the fluorescent column width from the mean vascular diameter obtained from brightfield images. 7. Divide the resulting value by 2 to obtain the thickness of the glycocalyx on the selected microvessel (see Note 9).

4

Notes 1. Catheters for rodent vessel cannulation can be commercially purchased ready to use or handmade from polyethylene 50 (PE50) tubing. Instructions on how to hand-make catheters are provided in a previously published protocol [23]. 2. The value of 2.0–2.5% is a suggestion for isoflurane maintenance. This value may vary by animal. Continue to monitor the animal’s breathing throughout the protocol and adjust as needed. 3. Consultation of a resource such as Anatomy and Dissection of the Rat third ed. by Walker and Homberger may be helpful for localizing the jugular vein [24]. 4. It is important to take the mesentery out by holding the intestinal walls. This prevents the delicate microvessels of the mesentery to break up and hemorrhage. 5. Intravital microscopy stages are commercially available. As an alternative, you can make your stage using plexiglass. 6. It is important to keep the mesentery moist throughout the protocol. 7. To recognize postcapillary venules, look for a bifurcation with convergent direction of blood flow. Arterioles have divergent flow direction at bifurcations. 8. White blood cells take up FITC-lectin. If the venule you select shows a great amount of slow-rolling or adhered leukocytes, choose a different area to analyze. Ideally, an area where the glycan concentration can be quantified by FITC-lectin bound to the endothelial surface, not to leukocytes, is desired. 9. This method assumes equal glycocalyx layer thickness on both walls of the microvessel.

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Acknowledgments The authors acknowledge their support from NIH grant R01GM120774. References 1. Reitsma S, Slaaf DW, Vink H et al (2007) The endothelial glycocalyx: composition, functions, and visualization. Pflugers Arch 454:345–359 2. Weinbaum S, Tarbell JM, Damiano ER (2007) The structure and function of the endothelial glycocalyx layer. Annu Rev Biomed Eng 9: 121–167 3. Becker BF, Chappell D, Jacob M (2010) Endothelial glycocalyx and coronary vascular permeability: the fringe benefit. Basic Res Cardiol 105:687–701 4. Yen WY, Cai B, Zeng M et al (2012) Quantification of the endothelial surface glycocalyx on rat and mouse blood vessels. Microvasc Res 83: 337–346 5. Alves NG, Trujillo AN, Breslin JW et al (2019) Sphingosine-1-phosphate reduces hemorrhagic shock and resuscitation-induced microvascular leakage by protecting endothelial mitochondrial integrity. Shock 52:423–433 6. Florian JA, Kosky JR, Ainslie K et al (2003) Heparan sulfate proteoglycan is a mechanosensor on endothelial cells. Circ Res 93:e136– e142 7. Yang Y, Schmidt EP (2013) The endothelial glycocalyx: an important regulator of the pulmonary vascular barrier. Tissue Barriers 1(1): e23494 8. Tarbell JM (2010) Shear stress and the endothelial transport barrier. Cardiovasc Res 87: 320–330 9. Torres Filho I, Torres LN, Sondeen JL et al (2013) In vivo evaluation of venular glycocalyx during hemorrhagic shock in rats using intravital microscopy. Microvasc Res 85:128–133 10. Torres LN, Sondeen JL, Ji L et al (2013) Evaluation of resuscitation fluids on endothelial glycocalyx, venular blood flow, and coagulation function after hemorrhagic shock in rats. J Trauma Acute Care Surg 75:759–766

11. Rahbar E, Cardenas JC, Baimukanova G et al (2015) Endothelial glycocalyx shedding and vascular permeability in severely injured trauma patients. J Transl Med 13:117 12. Dura´n WN, Sa´nchez FA, Breslin JW (2008) Microcirculatory exchange function. In: Tuma RF, Dura´n WN, Ley K (eds) Handbook of physiology: microcirculation. Academic Press/Elsevier, San Diego, CA, pp 81–124 13. Gao L, Lipowsky HH (2010) Composition of the endothelial glycocalyx and its relation to its thickness and diffusion of small solutes. Microvasc Res 80:394–401 14. Mulivor AW, Lipowsky HH (2002) Role of glycocalyx in leukocyte-endothelial cell adhesion. Am J Physiol Heart Circ Physiol 283: H1282–H1291 15. Childs EW, Tharakan B, Hunter FA et al (2007) Apoptotic signaling induces hyperpermeability following hemorrhagic shock. Am J Physiol Heart Circ Physiol 292:H3179– H3189 16. Kataoka H, Ushiyama A, Kawakami H et al (2016) Fluorescent imaging of endothelial glycocalyx layer with wheat germ agglutinin using intravital microscopy. Microsc Res Tech 79:31– 37 17. Tabuchi A, Mertens M, Kuppe H et al (2008) Intravital microscopy of the murine pulmonary microcirculation. J Appl Physiol (1985) 104: 338–346 18. Lipowsky HH, Gao L, Lescanic A (2011) Shedding of the endothelial glycocalyx in arterioles, capillaries, and venules and its effect on capillary hemodynamics during inflammation. Am J Physiol Heart Circ Physiol 301:H2235– H2245 19. Chappell D, Jacob M, Rehm M et al (2008) Heparinase selectively sheds heparan sulphate from the endothelial glycocalyx. Biol Chem 389:79–82

Microvascular Glycocalyx Quantification 20. Salmon AH, Ferguson JK, Burford JL et al (2012) Loss of the endothelial glycocalyx links albuminuria and vascular dysfunction. J Am Soc Nephrol 23:1339–1350 21. Vink H, Duling BR (1996) Identification of distinct luminal domains for macromolecules, erythrocytes, and leukocytes within mammalian capillaries. Circ Res 79:581–589 22. Henry CB, Duling BR (1999) Permeation of the luminal capillary glycocalyx is determined by hyaluronan. Am J Phys 277:H508–H514

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23. Doggett TM, Tur JJ, Alves NG et al (2018) Assessment of cardiovascular function and microvascular permeability in a conscious rat model of alcohol intoxication combined with hemorrhagic shock and resuscitation. Methods Mol Biol 1717:61–81 24. Walker WF, Homberger DG (1997) Anatomy and dissection of the rat. W. H. Freeman, New York

Chapter 14 Measurement of Blood-Brain Barrier Hyperpermeability Using Evans Blue Extravasation Assay O’lisa Yaa Waithe, Xu Peng, Ed W. Childs, and Binu Tharakan Abstract Blood-brain barrier (BBB) dysfunction and hyperpermeability have been implicated in a myriad of brain pathologies. The Evans Blue assay is one of the most popular methods for studying BBB integrity and permeability in rodent models of brain disorders. Under normal physiological conditions, the BBB is impermeable to albumin, so Evans Blue when injected intravenously binds to serum albumin and remains restricted within blood vessels. In traumatic and ischemic injuries, and other brain pathologies that result in BBB hyperpermeability, neighboring endothelial cells partially lose their close contacts to each other, and the BBB becomes permeable to proteins such as albumin. This paracellular leak of Evans blue-bound albumin is considered a reliable indicator of BBB dysfunction and hyperpermeability. Here, we describe the procedures for the evaluation of BBB integrity and hyperpermeability using Evans Blue extravasation assay in a mouse model of traumatic brain injury. The method described here focuses on intravenous injection of Evans Blue followed by Evans Blue dye extraction. This is followed by the measurement of fluorescence intensity of Evans Blue to determine the dye extravasation as a direct indicator of BBB hyperpermeability. Key words Blood-brain barrier, Microvascular hyperpermeability, Evans Blue assay

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Introduction The blood-brain barrier (BBB), the protective barrier of the brain, plays a critical role in controlling the influx and efflux of biological substances essential for the brain’s metabolic activity as well as neuronal function. Thus, the structural and functional integrity of the BBB is pivotal to maintain the homeostasis of the brain microenvironment [1–5]. The BBB is comprised of capillary endothelial cells alongside a basement membrane, accompanied by other cells such as pericytes and astrocytes. Structurally, the BBB consists of endothelial tight junctions, facilitated by tight junction proteins (TJPs). Tight junction proteins play an essential role in regulating the permeability across the blood-brain barrier [1–5, 8, 9]. Tight junction proteins can be, most often, integral transmembrane proteins that include claudins, occludin, and junctional adhesion

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molecules that are linked to tight junction-associated proteins, particularly to the accessory cytoplasmic membrane protein Zonula occludens-1 (ZO-1). The BBB is not a static structure, and in addition to TJP expression, its permeability can be affected by a multitude of factors; size, shape, charge, the type of the extravascular molecules, the nature of the vascular beds, etc. are all factors that contribute to barrier integrity and permeability. Furthermore, barrier permeability can also be affected by damage or injury conditions, or by other pathologies [6–15]. For instance, the vascular hyperpermeability observed due to of BBB dysfunction has been shown to be a significant cause of cerebral edema and elevated intracranial pressure following traumatic brain injury (TBI) [11–15]. Traumatic brain injury is a condition in which normal brain function is disrupted, either by external or internal force or trauma. This microvascular endothelial hyperpermeability observed at the level of the BBB remains a reliable marker for TBI for the reason that it is also a significant problem common in many other vascular conditions associated with traumatic, hemorrhagic, and ischemic injuries [1–4, 11–16]. Of the types of edema seen resultant from TBI, vasogenic edema is shown to be the most significant form occurring via BBB dysfunction and resulting hyperpermeability [1, 2]. The method detailed here first mentions the procedure of inducing TBI using a controlled cortical impactor (CCI). Though barrier permeability changes have been discovered to play a role in a multitude of pathologies, human brain research comes with many challenges and constraints; hence, there is a need for a reliable animal model for conditions producing such barrier changes [1, 2]. The CCI model of TBI described here uses a targeted mechanical impactor to replicate an external force inflicting injury to the mouse brain. Blood-brain barrier disruption is a hallmark feature of the secondary injury that occurs following TBI, frequently associated with leakage of fluid and proteins into the extravascular space leading to vasogenic edema and elevation of intracranial pressure [15–17]. This method offers the potential to make use of the Evans Blue extravasation assay as a reliable and easily doable approach to measure BBB dysfunctions and microvascular hyperpermeability following traumatic, ischemic, and other conditions of brain pathology. Evans Blue dye is a pigment that when injected into blood, binds to serum albumin. Due to the fact that under normal circumstances this albumin is too large of a molecule to pass the barrier, this bound albumin can then become an indicator for barrier dysfunction and microvascular hyperpermeability following TBI, based on concentrations observed on either side of the barrier. Evans Blue (EB) dye has owned a long history as a biological dye and diagnostic agent. It is an alkaline dye (MW- 961 daltons) that binds to serum albumin. In addition to its effective use in

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determining the integrity of the BBB in animal models of brain pathology, it has been used to assay changes in vascular permeability in various physiological disorders, including but not limited to immunological, inflammatory, cardiovascular, and cancer research. In rodent research, Evans Blue can be administered through various routes, such as intravenous, intraperitoneal, and subcutaneous, depending on the requirements of the study and the organ under investigation. Under normal conditions, there is a restricted exchange of fluids and solutes occurring at the blood-brain barrier. However, in conditions such as TBI, this restricted exchange is altered or impaired leading to BBB hyperpermeability. Evans Blue thus works under the principle that homeostatic conditions of the BBB showing minimal leakage of albumin into the brain, with a breach in barrier integrity leading to an observable leakage. Changes in BBB permeability using the dye measures can be either quantified by visualization, e.g., microscopy or by fluorometric/ colorimetric techniques.

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Materials

2.1 Reagents and Working Solutions

• Phosphate-buffered saline (PBS). • 70% alcohol. • Trichloroacetic acid. • 0.9% saline. • Betadine solution. • Anesthetic agent – urethane. • 5% Evans Blue solution.

2.2

Equipment

• Controlled cortical impactor (for inducing traumatic brain injury). • Cooling centrifuge. • Heating pad or lamp. • Electric razor. • Tissue homogenizer.

2.3

Supplies

• Sterile cotton swaps. • Gauze. • Cover glass with a diameter of 5 mM. • Lubricating eye ointment. • Syringes. • Needles (size: 25G, 27G/30G). • Gloves and other PPEs as required.

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Methods

3.1 Anesthetizing the Animals

Approval from an institutional animal care and use committee (IACUC) is required prior to starting animal studies. Additionally, it is advised to consult with institutional veterinarian prior to the start of any animal procedures. Male C57BL/6 mice (2–5 months old) are utilized in this study. Anesthesia type is normally based on the kind of surgery performed, the desired duration of anesthesia effect, route of administration (i.e., oral or IP), etc. Wakefulness of the animal should be examined prior to any procedures, typically by toe-pinch or tail-pinch response. Following anesthesia, eye-lubricating ointment is applied to prevent dryness of the eyes. To ensure the animal maintains proper body temperature, it must be placed under a lamp or on a circulating water heat pad throughout the duration of the procedure. A rectal probe can be used to monitor the body temperature of the animal, if required. In this study, urethane is used as a terminal anesthesia in mice. The animal should be warmed sufficiently prior to the urethane injection (via either heat pad or lamp as stated before), as well as for the duration of the procedure. Urethane is found to be the optimal choice for these experiments because it can maintain effects at a constant level for up to several hours.

3.2 Intravenous Injection of Evans Blue

In this protocol, Evans Blue dye was injected intravenously via the tail vein, with the reason being that it is noninvasive as compared to catheters and the related procedures. Care should be taken to inject the dye into the lateral tail vein. Sterility of the injected dye/solutions and the workplace must be maintained throughout the procedure, as they can affect animal mortality rates. If necessary, veins can be dilated for further ease of procedure via warming the mouse either using a heat lamp, heat pad, or immersing the tail in warm water. To ensure that the dye is properly heated to the body temperature of the animal, prior to injection, let it sit in a warm water bath (around 37–38 °C). Standard tail vein injection volumes for mice do not typically exceed 200–250 μL, though the exact maximum volume may depend on the size, strain, and vehicle used.

3.3 Surgical Procedures

To avoid dehydration, animals should be injected with 1 mL isotonic saline solution. Body temperature of 37–38 °C must be always maintained, via use of a heating pad or lamp. This temperature can be checked via use of a rectal probe. The following are the step-by step surgical procedures for inflicting traumatic brain injury in mice: 1. Shave the dorsal side of the animal’s head using an electric razor (alternatively, a hair remover gel may be used).

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2. Wipe the area with gauze or sponge soaked in ethanol (70%), followed by application of betadine solution at the site. 3. Apply the eye lubricant directly to the eyes of the animal using a sterile cotton swab. 4. Inject Evans Blue dye as described above. 5. Animals are then prepped for either sham or traumatic injury procedures for CCI. 6. The methods described here utilize the benchmark stereotaxic impactor from Leica. Dwell time, velocity, and impactor tip size should be decided and adjusted beforehand to determine the severity of trauma. Impactor position is determined by the placement of the animal’s head. 3.4 Traumatic Brain Injury

Begin the sham injury or traumatic injury as required: 1. Place the extend/retract toggle switch into the “extend” position. 2. Sham animals are then mounted on the stereotaxic frame: hold the animal toward one side of the bar. Push the other side into the ear canal and tighten after putting the fixture. 3. Center the impact probe onto the lambda position and the anterior/posterior, medial/lateral readings, and zero the digital manipulator. 4. The clip located at the end of the contact sensor’s lead wire should be pinned to the skin, ear, or tail base. Impact probe is lowered onto cranium until a beep is heard. 5. The dorsal/ventral reading on digital manipulator is zeroes, and the retract/extend toggle switch is flipped once again to the “retract” position. 6. Dorsal/ventral settings on the digital manipulator can be adjusted to negative values based on the injury depth chosen for the study (e.g., 1 was chosen for mild TBI, -2 for moderate, and -3 for severe injury). 7. Animal is then removed from the platform and placed in an incubator for recovery and proper monitoring prior to placement back in cage or to another pad/lamp for euthanasia.

3.5 Evans Blue Injection

Following anesthesia, each animal is injected with the Evans Blue dye (5% concentration, intravenous, up to 100 μL). Prior to injection, the dye can be dissolved in sterile PBS/0.9% saline. Tail vein injection is performed as described in the IV injection protocol section. Evans Blue should be injected 5–30 min prior to procedure.

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3.6 Transcardial Perfusion and Tissue Collection

Upon reaching the endpoint of the study, animals are perfused transcardially with ice-cold sterile saline with heparin (1000 U/ mL). This ensures the removal of any blood clots present (at least 25 mL). The perfusion removes the intravascular Evans Blue (EB) dye, leaving the remaining EB dye in extravascular space of the brain tissue. The animal is then decapitated, and the brain and any other organs of interest are harvested. Harvested brains can be used for trichloroacetic (TCA) assay and can be used to extract the dye followed by the assay immediately or stored at -80 °C for later experiments as required.

3.7 Evans Blue Dye Extraction

Brain hemispheres are first homogenized using a tissue homogenizer in 50% TCA (w/v). The volume of TCA chosen is adjustable based on weight of the select brain tissue taken. Subsequent homogenate will be centrifuged at 10,000 rpm for 20 min. Supernatant collected will is diluted in three parts ethanol (1:3, 50% TCA-ethanol). Evans Blue dye can be quantitated either by fluorimetry (620 nM emission/680 nm excitation) or colorimetry (610 or 620 nM) techniques. Samples are finally evaluated via corresponding standards ranging from 50 to 1000 ng/mL (Fig. 1).

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Notes 1. Animals can either be anesthetized in several ways using a combination of methods or by individual substances such as isoflurane (4% for induction and 1.5–2% for surgery; with 40% O2 and 60% N2 via use of a facemask) or urethane as described in this procedure. 2. Warming lamps provide risk of burn injury and thus should be kept at safe working distance. 3. Animals should always have a gauze or cloth blanket separating them from the warming pad – they should never be laid directly atop the pad. Continuously monitor the animal for burns. 4. Urethane is carcinogenic and used for terminal anesthesia; hence, it should be handled carefully and with discretion. Steps should be taken by the researcher preparing the procedure, always including the use of personal protective equipment (PPE). Urethane should only be prepared and injected to animals under a chemical hood. 5. Warm water should be handled around the animal with caution. High-temperature water exposure for too long can scald the tail tissue leading to a burn injury. 6. EBD is not considered toxic; however, it is still a hazardous irritant and a potential carcinogen. Researchers should take caution in handling.

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Fig. 1 This figure details the extravasation of Evans Blue in the brain of a mouse model of traumatic brain injury, demonstrating BBB hyperpermeability. Post-CCI brains were harvested for Evans Blue analysis. The data shows that melatonin pretreatment attenuates TBI-induced BBB hyperpermeability significantly (Panel a; p < 0.05). Panel b shows the representative images of the brain tissue obtained from various experimental groups. Melatonin (10 μg/g body weight) pretreatment significantly decreased TBI-induced Evans Blue extravasation (p < 0.05). Mice were divided into a sham group (n = 6), a vehicle + sham group (n = 6), a vehicle + TBI group (n = 5), and a melatonin + TBI group (n = 6). Data represents ng/brain cortex ± SEM. “*” indicates statistical significance. “a” indicates significant increase compared to the sham injury group or vehicle + sham injury group. “b” indicates significant decrease compared to vehicle + TBI group. This figure is taken from our previously published study. (Alluri et al, Plos One. 2016 May 6;11 [5]:e0154427). Creative Commons License https://creativecommons.org/licenses/)

7. Perfusion protocol: The animal is placed on its back and the limbs are pinned. Forceps are used to lift the skin and body wall. An incision is made laterally, followed by two additional ones on either side through the ribcage. The diaphragm is then cut. The sternum is lifted. The loose flap is then pinned and a

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needle containing saline is inserted into the left ventricle. A small incision is made in the right atrium. Animal is perfused for 10 min (or until all the blood is perfused out from the right atrium). The animal is then decapitated via guillotine and tissues are harvested. 8. Repeat experiment multiple times to ensure validity.

Acknowledgments The authors acknowledge the support from National Institutes of Health (NIH) grant 5 SC3 NS127765-02 (BT). References 1. Alluri H, Wiggins-Dohlvik K, Davis ML, Huang JH, Tharakan B (2015) Blood-brain barrier dysfunction following traumatic brain injury. Metab Brain Dis 30(5):1093–1104 2. Alluri H, Anasooya Shaji C, Davis ML, Tharakan B (2018) A mouse controlled cortical impact model of traumatic brain injury for studying blood-brain barrier dysfunctions. Methods Mol Biol 1717:37–52 3. Kumar P, Shen Q, Pivetti CD, Lee ES, Wu MH, Yuan SY (2009) Molecular mechanisms of endothelial hyperpermeability: implications in inflammation. Expert Rev Mol Med 11:e19 4. Marques PE, Oliveira AG, Amaral SS, NunesSilva A, Almeida AFS (2012) Intravital microscopy: taking a close look inside the living organisms. Afr J Microbiol Res 6:1603–1614 5. Shen Q, Wu MH, Yuan SY (2009) Endothelial contractile cytoskeleton and microvascular permeability. Cell Health Cytoskelet 1:43–50 6. Masedunskas A, Porat-Shliom N, Tora M, Milberg O, Weigert R (2013) Intravital microscopy for imaging subcellular structures in live mice expressing fluorescent proteins. J Vis Exp 79:50558 7. Nidavani RB, Mahalakshmi AM, Shalawadi M (2014) Vascular permeability and Evans blue dye: a physiological and pharmacological approach. J Appl Pharm Sci 4:106–113 8. Yuan SY (2002) Protein kinase signaling in the modulation of microvascular permeability. Vasc Pharmacol 39:213–223

9. Radu M, Chernoff J (2013) An in vivo assay to test blood vessel permeability. J Vis Exp 73: e50062 10. Deaglio S, Robson SC (2011) Ectonucleotidases as regulators of purinergic signaling in thrombosis, inflammation, and immunity. Adv Pharmacol 61:301–332 11. O’Connor WT, Smyth A, Gilchrist MD (2011) Animal models of traumatic brain injury: a critical evaluation. Pharmacol Ther 130:106–113 12. Parikh S, Koch M, Narayan RK (2007) Traumatic brain injury. Int Anesthesiol Clin 45: 119–135 13. Unterberg AW, Stover J, Kress B, Kiening KL (2004) Edema and brain trauma. Neuroscience 129:1021–1029 14. Xiong Y, Mahmood A, Chopp M (2013) Animal models of traumatic brain injury. Nat Rev Neurosci 14:128–142 15. Gean AD, Fischbein NJ (2010) Head trauma. Neuroimaging Clin N Am 20:527–556 16. Kasper C, Yvette C (2015) Traumatic brain injury. Annu Rev Nurs Res 33:xi-xii 17. Khan M, Im YB, Shunmugavel A, Gilg AG, Dhindsa RK, Singh AK, Singh IJ (2009) Administration of S-nitrosoglutathione after traumatic brain injury protects the neurovascular unit and reduces secondary injury in a rat model of controlled cortical impact. Neuroinflammation 6:32

Chapter 15 Assessment of Endothelial Barrier Functions in Extra Embryonic Vasculature of Chick Embryo as an Alternative Model Jamila Siamwala, Akila Swaminathan, and Suvro Chatterjee Abstract Vascular permeability, a tightly regulated process, is a direct measure of angiogenic and immune responses in the endothelium altered in several acute and chronic diseases such as sepsis, high-altitude pulmonary edema (HAPE), high-altitude cerebral edema (HACE), ischemia, and coronavirus disease 2019 (COVID19) endotheliitis. Both endogenous and exogenous factors such as cytokines, chemokines, and hormones may affect vascular permeability. The conventional tools available for the measurement of vascular permeability in vitro and in vivo based on collagen-coated Transwell and dye-based spectrophotometric methods are indirect measures of permeability. In this chapter, we present our live in ovo protocols based on dextranTexas red and avian chorioallantoic membrane assay developed using custom-made equipment to assess leakiness of endothelial cell barrier both in vitro and in vivo. Further, we validate this assay using different stressors such as ischemia and hypoxia known to affect endothelial barrier properties by potentiating actin stress fiber disorganization and disrupting the cell-cell junctions. Key words Endothelial barrier functions, Angiogenesis, Immune functions, Chorioallantoic membrane, COVID-19 endotheliitis, Sepsis, Hypoxia, Ischemia, Edema

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Introduction Vascular permeability is an essential element of angiogenesis [1], inflammation [2], leukocyte extravasation [3], and tissue damage in most of the organ diseases. Sepsis [4], high-altitude pulmonary edema (HAPE) [5], high-altitude cerebral edema (HACE), and COVID-19 endotheliitis [6–8] are morbid diseases associated with increased vascular permeability, multi-organ failure, and death. In severe sepsis, inflammatory mediators chronically activate the endothelium leading to increase in vascular permeability and leakage of plasma components and pathogens in the affected tissue subsequently resulting in organ failure. As a result, patient with sepsis typically develop subcutaneous and body cavity edema associated with the accumulation parenchymal and interstitial fluid

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through the damaged vasculature [9]. Recovery from acute septic shock requires restoration of vascular permeability and reduction in edema. Vascular permeability and edema are critical prognostic determinants of the debilitating, inflammatory sepsis condition [10]. Apart from sepsis, high-altitude pulmonary edema (HAPE) and high-altitude cerebral edema (HACE) are two potentially lifethreatening pathological conditions that occur due to hypoxic vasoconstriction and endothelial dysfunction occurring at altitudes greater than 3000 m5. Endothelial remodeling is primarily associated with monolayer leakiness at hypoxic conditions. The imbalance in the alterations in intermediators of vascular tone and morphological changes in the endothelial monolayer play an important role in hypoxia-mediated pathogenesis. Therefore, hypoxia is another critical inducer of vascular permeability in pulmonary diseases [11–13]. Recent postmortem histology evaluation of patients with coronavirus disease caused by severe acute respiratory syndrome coronavirus 2 (SARS CoV-2) showed lymphocytic endotheliitis of the liver, lung, heart, kidney, and submucosal vessels of the small intestine. COVID-19 endotheliitis is suggested to be the primary cause of systemic microcirculatory dysfunction in the multiorgan vascular beds and sequelae of lung injury, pathological angiogenesis, and endothelial dysfunction-associated apoptosis in COVID-19 patients [8]. Pathological angiogenesis is a hallmark of ischemic diseases such as myocardial infarction and stroke and occurs as a compensatory mechanism to reperfuse ischemic regions. The microvasculature consisting of arterioles, capillary network, and postcapillary venules primarily control the vascular tone and vascular permeability and is indispensable for vascular homeostasis [14]. Endothelial dysfunction is a hallmark of microvascular dysfunction specifically vasoconstriction and subsequent organ ischemia, inflammation, and procoagulant state [15]. Endothelial cells form a semipermeable barrier between blood and tissue mediating paracellular transport of gas, solutes, nutritional factors, and circulating hormones. Endothelial monolayer interfaces between blood and the vessel wall transporting blood-borne molecules into the tissues and releasing cell secretome into the blood stream, thereby contributing to the regulation of vascular homeostasis. The integrity of the barrier is site specific and varies between the organs and within the same vasculature, for example, in the lungs. The mechanisms involved in endothelial cell barrier integrity include vascular endothelialcadherin (VE-cadherin, CD144) [16]. Pro-inflammatory cytokines, chemokines, and reactive oxygen species may induce apoptosis [17] and pyroptosis [18] compromising vascular integrity and leading to capillary leak and entry of vascular contents in the tissue and extracellular space. Perturbations in the vascular bed may cause

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endothelial monolayer remodeling affecting the vascular tone. Hypoxia, another endothelial cell activator, further exaggerates vasoconstriction, leading to capillary stress failure. Upon endothelial cell activation by hypoxia, the actin cytoskeleton disorganizes resulting in inter-endothelial gaps followed by enhanced paracellular endothelial permeability [19]. The actin cytoskeleton organization and adherens junction proteins are regulated by Rac and Rho family of GTPases [20–22 ]. The small GTPase RhoA increases endothelial cell permeability in response to edematous agents such as a-thrombin and histamine by modifying actin stress fibers [23], focal adhesion complexes (FAC), adherens junction protein, tight junction proteins, and F-actin fibers [24]. In addition, RhoA mediates Protein Kinase C (PKCd)-induced protection of endothelial cell barrier dysfunction and reorganization of focal adhesion kinases and F-actin fibers [25]. Further, S1P2R-Rho-ROCK-PTEN axis regulates endothelial cell paracellular permeability [26]. Cigarette smoke causes pulmonary edema by disrupting the endothelial cell barrier via oxidative-mediated inhibition of RhoA and FAC [27]. In summary, endothelial cell permeability is a prima facie determinant of vascular homeostasis and vascular dysfunction. Therefore, development of analytical tools to study live endothelial cell permeability in vitro and in vivo is a prerequisite in cell biology and related biomedical research disciplines. Limited indirect and qualitative approaches are available to track single-cell activity in hypoxic conditions lacking in sensitivity and accuracy. The current tools available for the measurement of vascular permeability in vitro and in vivo based on collagen-coated Transwell and dye-based spectrophotometric methods are indirect measurements. There is an urgent need to develop novel approaches to facilitate live cell permeability tracking at single cell, cell monolayer, and organ level. Commonly used in vitro methods of endothelial cell permeability such as two-compartment Transwell chamber and electrical cell impedance sensor (ECIS) technique are based on indirect measurements of endothelial cell permeability. The two-compartment Transwell chamber consists of 0.4 μM pore polycarbonate Transwell constructs on which the endothelial cells are seeded. The endothelial cells are then treated with permeabilizing compounds or inhibitors of permeability. Accumulation of the compounds in the lower chamber is determined using spectrophotometric assays [28]. ECIS method of permeability detection is based on the measurements of electrical resistance across confluence endothelial cell monolayers. In this method, endothelial cells are coated on collagen-coated gold electrodes, and the electrical resistance is recorded when the impedance reaches ≥500 ohms [29]. The disadvantage of these techniques is that they are costly and indirect and lack shear stress and flow which are the part of circulation. We developed single-cell trypan blue inclusion method

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measuring the rate of inclusion using microscopic photometry principles. This is based on the principle that healthy cells exclude trypan blue compared to dead cells which retain the trypan blue dye. In vivo methods include high-resolution intravital microscopy used to detect cells involved in paracellular transport and hemophilic and heterophilic cell-cell interaction in rodents. However, the specialized skills, equipment, time, and complicated analysis are required to perform such analysis. Moreover, adherences to the 3Rs (replacement, reduction, and refinement) are required to conduct animal research which further restricts the development of any new technique. To circumvent these problems, we have developed a simple device to image live vascular permeability using chorioallantoic membrane stressed with hypoxia. Chorioallantoic membrane is used to track vascular development in an open fertilized egg incubated at 37 °C. Based on the morphological landmarks, the chick embryos are staged according to Hamburger-Hamilton (HH) stages [30]. We used the chick embryo stage 12–13 HH CAM possessing 16 somites formed by progressive segmentation of the paraxial mesoderm, head-fold and neural folds for the development of the permeability assay. Previously, we have developed ischemia CAM model by ligating the right vitelline artery using sterile surgical suture using the same stage 12–13 embryo [31]. The advantages of using CAM apart from adhering to the three Rs (replacement, reduction, and refinement) are to allow realtime monitoring of vascular permeability in its native physiological condition in the presence flow and shear stress. The method is elaborated in the Materials and Methods section. Further, we show endothelial cell remodeling using inducers of vascular permeability such as cadmium, sFRP4, and inhibitors of endothelial cell permeability such as nitric oxide and cGMP analog 8-Br cGMP. Cadmium, a potent toxicant in cigarette smoke, causes endothelial cell permeability associated with myocardial infarction, hypertension, and atherosclerosis [32]. We show via endothelial permeability experiments that nitric oxide donor, spermine NONOate, rescues cadmium-mediated endothelial cell permeability [33]. Secreted Frizzled-Related Protein 4 (sFRP4) mediates its anti-angiogenic functions by targeting nitric oxide in the endothelium [34]. Our experiments indicate that nitric oxide and cGMP analog 8-Br cGMP may further reduce sFRP4-mediated increases in endothelial cell permeability [34]. In another experiment, using the new technology, we show that in addition to reducing blood pressure, dietary nitrates derived from fennel seeds promote vascular homeostasis and protect the endothelial cell monolayer from hypoxia-induced leakiness. In summary, we have developed a technique to facilitate real-time high-resolution tracking of Texas Red across the vascular wall of chorioallantoic membrane in presence of inducers or inhibitors of endothelial cell permeability in addition to

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disease models of hypoxia and ischemia. The platform enables (1) reproducible, quantitative measurement of angiogenic responses, (2) simple and reliable testing of endothelial cell permeability mechanisms in simulated disease conditions, (3) visualization of multiple processes during shear stress and flow in homeostasis and disease, (4) multiplexing of drugs and compounds that affect endothelial cell permeability, and (5) industrial uses as a costeffective, rapid drug screening tool and cutting-edge mechanics like manipulation fluidics. Moreover, the CAM provides holistic information on the vasculature comprising of arteries, veins, and capillaries in a physiological setting. The CAM is immunodeficient, develops rapidly, and is easily accessible. In addition, there are fewer ethical concerns using CAM as a disease model. Further research using appropriate image processing platform for the images or video acquired by this technique would enable generation of high precision data for the vascular barrier functions. 1.1

Limitations

The main limitation of the assay is that not all the vascular outgrowths and angiogenesis can be accounted for as this assay is performed only on the visible vascular structures in the camera. Secondly, there is a critical window of drug application which enables the measurements of vasodilation and permeability. This can be overcome by video recording for longer periods or by timelapse imaging. The CAM is also sensitive to pH, temperature, oxygen tension, and salinity, and therefore resealing after breaking the eggs is critical. There are reports on inflammation-induced proliferation of blood vessels in addition to the drug and the primary effect. Reducing the time of the procedure and minimal disturbance during embryo handing and appropriate controls would minimize the nonspecific effects. Further embryo survival rate is low and the embryos may not reach hatching. In conclusion, CAM model of vascular permeability is a simple, reliable, reproducible, and quantitative model to perform angiogenic, immune, and vascular permeability assessment of the entire embryonic cardiovascular system with no ethical concerns. The present model allows testing the multiple compounds in physiological and disease conditions and is cost-effective and less complex. This model reduces use of animals in experimental research and supports the 3R rules. The limitation of the present model is time duration of the “viable phase” of the model which is restricted to a maximum of 8 h. The model has diagnostic, mechanistic, and therapeutic applications in vascular research and pharmaceutical investigation.

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Materials Instrumentation

(i) Hypoxia Chamber. 1. Hypoxia chamber consists of an inlet, an outlet, inner space, and airlock system. Inlet is used for purging the hypoxia gas (5% oxygen and 95% nitrogen) into the chamber. 2. The outlet is for the release of the gas after purging the hypoxia gas on the chick vascular bed, and the inner space/ closed chamber holds the testing plate, chick embryo, and premix gas. The airlock system prevents the outside air. 3. The hypoxia chamber is kept in the 37 °C incubator to maintain the temperature. (ii) Camera Setup for the In Ovo Assay. To capture images, the Olympus IX71 epifluorescence microscope is equipped with DP71 Camera (Olympus). The images are captured at 4× magnification.

2.2 Fennel Seed Extract (FSE) Preparation

1. Soak 3 g of fennel seeds in 20 mL of sterile distilled water and store at room temperature (25 °C) overnight. The overnight incubation is required for extraction (simple water extraction) from the fennel seeds. 2. On the next day, filter the extract using Whatman No. 1 paper. The Whatman No. 1 paper is used to separate the water extracts from fennel seeds. A tenfold dilution of FSE extract (effective concentration) was used for experimentation.

2.3 Texas Red Preparation

1. Texas Red conjugated with bovine serum albumin labels proteins and enables visualization of the protein through the blood vessels. Prepare the stock solution of Texas Red by dissolving 5 mg of Texas Red in 1× PBS with pH of 7.4. Wrap a foil around the tube to keep the Texas Red solution in the dark. The final working concentration of 10 μM Texas Red is prepared dissolving in PBS. Texas Red probe excitation is 596 and emission is -615.

2.4 Cell Culture Medium

Dulbecco’s Modified Eagle’s Medium (DMEM) with 10% fetal bovine serum (FBS) and 1% penicillin-streptomycin.

2.5 Materials Required for CAM Experiments

1. Fertilized brown leghorn chicken eggs, glass bowl, distilled water, 1× PBS, sterile scissors, forceps, and sharp knife. Sterilize Petri plates and 0.5 mm surgical suture. 2. Hematoxylin and eosin stain, 10% formalin, 0.1% Triton X. Prepare 0.1% of Triton X by dissolving 10 μL of Triton X in 10 mL of 1× PBS. Prepare 10 μM of Alexa Fluro Phalloidin from a stock solution of 10 mM.

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1. Dissolve 0.2 g of paraformaldehyde in 8 mL of 1× PBS. 2. Next, add 50 μL of 1 M NaOH into the paraformaldehyde solution (partially dissolved) and heat the solution at 90 °C for 10 min. 3. Once the solution cools down, adjust the pH to 7.0. Make up the volume to 10 mL with water. 4. Finally, the solution made up to 10 mL.

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3.1 In Vivo Chorioallantoic Membrane Permeability Assay Using Texas Red

1. Incubate the 4-day fertilized eggs (Poultry Research Station, Potheri, Chennai) at a 37 °C incubator. Texas Red permeability assay is performed on CAM treated with control (DMEM Medium) and FSE. Incubate the CAM in normoxia and hypoxia conditions for 1 h at 37 °C [21, 22] (Figs. 1, 2, 3, and 4). 2. Break the eggs gently and carefully make a small hole on the lower portion of the broader end of the egg and gently remove as much albumin as possible (Fig. 1). 3. The egg shell was removed carefully without disturbing the yolk/embryo and transfer contents to a sterile Petri dish immersed in distilled water. 4. The egg yolk was turned gently so the embryo was visualized.

Fig. 1 Preparation of shell-less chick embryo culture. (a) Wipe the eggshells with 70% alcohol solution to sterilize and remove impurities. (b) Make a tiny hole at the narrow end of the eggshell with a board pin. (c) Remove excess of albumin with the 1 mL pipette to prevent overflow while opening the broader end of the egg. (d) Seal the hole with adhesive tape to prevent leakage of the albumin and yolk. (e) Remove the eggshell from the broader end of the egg with forceps without the disturbing the embryo. (f) Transfer the entire contents into the sterile dish. (g) Representative image of in vivo chick vitelline membrane visualized in the camera fixed on top of the phase contrast microscope. Yellow-colored arrow indicates the boundary of the membrane which prevents dye permeation. Blue-colored arrow indicates the dark hallow inside the blood vessel

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Fig. 2 Validation of the model. (a) Hypoxia effects on endothelial integrity on chick vascular bed. (A) The vascular bed was treated under normoxic condition for 2 h. The membrane was stained with Alexa Fluor phalloidin (10 μM). (B) The expanded view of blood capillaries. This figure shows the intact monolayer of cells observed under normoxic condition. (C) The vascular bed treated under global hypoxic condition for 2 h. (D) The white arrow indicates that blood capillaries showed more number of paracellular gaps (n = 3). (E) The vascular bed was treated under normoxic condition for 2 h. (F) This figure depicts that there are no gaps formed under normoxic condition. (G) The vascular bed was treated under ischemic condition for 2 h. (H) The yellow arrow indicates that ischemic treated blood capillaries formed more paracellular gaps compared to normoxia (n = 3). (b) The chick embryo model. The chick vascular bed contains different arteries and veins which are denoted as A –(anterior vitelline vein), B –(right vitelline artery), C –(right lateral vitelline vein), D – (posterior vitelline vein), E –(left vitelline artery), and F –(left lateral vitelline vein. (i) The hypoxic chamber contains inlet, outlet, and airlock system. The hypoxic gas mixture was perfused through the inlet. The shellless embryo was transferred into DMEM/HEPES contained in Petri dishes. Then, the Petri dish was kept inside the hypoxic chamber at 37 °C. (ii) The schematic representative images of Texas Red permeability assay in chick vascular bed. (iii) The most responsive blood vessel is the right vitelline artery. This section shows the thin layer of smooth muscle cells around the arteries. The epithelial cells and endothelial cells are compactly arranged in the vessels. Scale bar is 50 μM and magnification is 20×. Arv, area vasculosa; Lbv, large blood vessels; Sbv, small blood vessel; Ec l, ectodermal layer; En l, endodermal layer; V, vein; ep, epithelial cells; Bi, blood islands or blood sinus. The arrowhead indicates the presence of a smooth muscle layer around the vessels

5. Now, the CAM was detached from egg yolk using sharp scissors and forceps (see Note 1). 6. The membrane was scooped gently into a small Petri dish and carefully pipette out the water without disturbing the CAM. 7. The CAMs were gently washed with 1× PBS to remove excess yolk and albumin (see Note 2). 8. The CAMs were treated with FSE and control under normoxia and hypoxia conditions.

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Fig. 3 Summary of the in vitro and in ovo protocols developed to measure permeability. (i) In vitro single cell trypan blue inclusion method to measure endothelial cell permeability. (ii) Trypan blue exclusion method developed to measure permeability in endothelial cell monolayers. (iii) In ovo measurement of vascular permeability in CAM using Texas Red assay

9. Following treatment the CAMs were again gently washed with 1× PBS before being incubated with 10 μM of Texas Red for 30 min. 10. After treatment the CAMs were washed with 1× PBS. Finally, the CAMs were fixed with 2% paraformaldehyde for 10 min. 11. After 10 min, the fixative was removed and CAMs were washed again with 1× PBS. 12. The fluorescent images were taken at 4× magnification using an Olympus 1X71 epifluorescence microscope equipped with DP71 Camera.

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A

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Normoxia 4 3.5 3 2.5 2 1.5 1 0.5 0

Hypoxia

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Fig. 4 FSE Protects hypoxia-induced endothelial leakiness. (a) The chick vascular bed treated under normoxic and hypoxic condition with FSE and without FSE. The result of the Texas Red permeability assay showed the extent of dye permeation (vessel outline depicted with white arrows) in normoxia, hypoxia, FSE, and hypoxia with FSE-treated blood vessels. This figure indicates that hypoxia-induced leakiness was protected by FSE. (b) Graphical representation of quantification of dye accumulation in the blood vessel, and the result indicates that the hypoxia-treated blood vessel showed more Texas Red dye permeabilization compared to normoxic-treated blood vessels. FSE with hypoxia-treated blood vessel significantly protects the dye permeabilization compared to hypoxia-treated control group blood. **P = 0. 001 vs normoxia, #P = 0. 001 vs hypoxia. Values represent means for each group ± SEM, one-way ANOVA, and LSD, n = 3 3.2 Creating Ischemic on Chick Vascular Bed

1. The 4-day fertilized eggs were opened in sterile dishes. 2. The embryo viability and blood flow were checked under the stereomicroscope. 3. The ischemic condition was created on chick vascular bed [19]. 4. Briefly, after 4 days of incubation, eggshells were carefully broken by gently hitting the shell in the middle of the egg with a sharp knife.

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5. Then the entire contents of the egg were transferred to a sterile glass bowl. 6. The glass bowls with the egg contents were covered with sterile glass lids and placed in incubator; the eggs were opened carefully without disturbing the egg yolk. 7. However, damaged eggs with ruptured vascular bed or with any visible abnormalities were not used for this study. 8. The chick right vitelline artery was blocked using surgical suture (see Note 3). 9. Then, glass bowls were incubated at 37 °C for 2 h. For control, the eggs were opened in sterile glass bowl and incubated at 37 ° C for 2 h. 3.3 Global Hypoxia Induction on Chick Vascular Bed

1. On the fourth day of incubation, eggshells were carefully broken by gently hitting the shell in the middle of the egg with a sharp knife. 2. Then the entire contents of the egg were transferred to a sterile glass bowl. 3. The glass bowl was kept in the hypoxia chamber (gas mixture: 5% O2 + 95% N2) (Fig. 2). 4. Then the hypoxia chamber was kept at 37 °C for 2 h. 5. For normoxia condition, the vascular beds were incubated at 37 °C, 5% CO2 for 2 h [28].

3.4 Phalloidin Staining in Chick Vascular Bed

1. The 4-day fertilized egg was opened in sterile Petri dishes. 2. The chick vascular bed was detached from the egg yolk using scissors. 3. The vascular beds were washed with 1× PBS to removes excess yolk. 4. Then the chick vascular beds were treated with global hypoxia, ischemic condition, as well as control for 2 h. 5. After 2 h, the chick vascular bed was washed with 1× PBS. 6. The vascular beds were fixed with 2% of paraformaldehyde for 10 min. 7. Next, the vascular beds were washed with 1× PBS and permeabilized with 0.1% of Triton X. 8. Then, the vascular bed was stained with Alexa Fluor-phalloidin (10 μM) and incubated for 10 min. 9. The vascular beds were washed with 1× PBS gently. 10. The fluorescent images were taken at 4× magnification using an Olympus 1X71 epifluorescence microscope equipped with DP71 Camera [26] (Fig. 2).

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3.5 Histology of the Vessel Middle of the Bed (VMD)

1. The 4-day fertilized eggs were opened in sterile dishes. 2. The vascular bed was detached from the egg yolk. 3. The most responsive location of the blood vessels is VMD of the right vitelline artery. 4. VMD area was fixed with neutral formalin. 5. Then, the membranes were dehydrated and embedded in paraffin. 6. The membrane section was stained with hematoxylin and eosin. 7. Histological images were taken at 20× magnification using an Olympus IX71 epifluorescence microscope equipped with DP71 Camera [27].

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Notes 1. Cut the membrane all way around and avoid accidently cutting blood vessels. 2. Ensure membranes was spread across the plate and maintain this while washing. 3. Ischemic condition was created in the right vitelline artery without disturbing the egg yolk and without rupturing the membrane.

Acknowledgments The authors acknowledge the Rhode Island Foundation grant (20190594), Alzheimer administrative supplement to COBRE CPVB (P20 GM103652), and TEAM UTRA grant from Brown University to JHS. References 1. Fukumura D, Gohongi T, Kadambi A, Izumi Y, Ang J, Yun CO, Buerk DG, Huang PL, Jain RK (2001) Predominant role of endothelial nitric oxide synthase in vascular endothelial growth factor-induced angiogenesis and vascular permeability. Proc Natl Acad Sci U S A 98:2604–2609 2. Uilkhelm DL (1968) [Increased vascular permeability in acute inflammation]. Patol Fiziol Eksp Ter. 12:3–16 3. Wiener J, Lattes RG, Pearl JS (1969) Vascular permeability and leukocyte emigration in allograft rejection. Am J Pathol 55:295–327 4. Wu R, Song X, Xu Y, Meng X (2000) Apoptosis of endothelial cells in alteration of microvascular permeability in lung during sepsis. Zhonghua Wai Ke Za Zhi 38:385–387

5. Manohar M (1979) What causes the microvascular permeability change in high altitude pulmonary edema? Circ Res 44:873–874 6. Fox SE, Lameira FS, Rinker EB, Vander Heide RS (2020) Cardiac Endotheliitis and multisystem inflammatory syndrome after COVID-19. Ann Intern Med 173:1025 7. Mosleh W, Chen K, Pfau SE, Vashist A (2020) Endotheliitis and endothelial dysfunction in patients with COVID-19: its role in thrombosis and adverse outcomes. J Clin Med 9 Z, Flammer AJ, Steiger P, 8. Varga Haberecker M, Andermatt R, Zinkernagel AS, Mehra MR, Schuepbach RA, Ruschitzka F, Moch H (2020) Endothelial cell infection and endotheliitis in COVID-19. Lancet 395:1417– 1418

Chick Embryonic Vasculature as a model of Endothelial Barrier Functions 9. Groeneveld AB, Tacx AN, Bossink AW, van Mierlo GJ, Hack CE (2003) Circulating inflammatory mediators predict shock and mortality in febrile patients with microbial infection. Clin Immunol 106:106–115 10. Hotchkiss RS, Moldawer LL, Opal SM, Reinhart K, Turnbull IR, Vincent JL (2016) Sepsis and septic shock. Nat Rev Dis Primers 2:16045 11. Ebeigbe AB (1984) Vascular membrane permeability during hypoxia. Pharmacol Res Commun 16:351–358 12. Hu DL, Yu YX, Liang R, Zhou SY, Duan SL, Jiang ZY, Meng CY, Jiang W, Wang H, Sun YX, Fang LS. (2019) [Regulation of hypoxia inducible factor-1alpha on permeability of vascular endothelial cells and the mechanism]. Zhonghua Shao Shang Za Zhi 35:209–217 13. Siggaard-Andersen J, Petersen FB, Hansen TI, Mellemgaard K (1969) Vascular permeability and plasma volume changes during hypoxia and carbon monoxide exposure. Angiology 20:356–358 14. Flammer AJ, Anderson T, Celermajer DS, Creager MA, Deanfield J, Ganz P, Hamburg NM, Luscher TF, Shechter M, Taddei S, Vita JA, Lerman A (2012) The assessment of endothelial function: from research into clinical practice. Circulation 126:753–767 15. Bonetti PO, Lerman LO, Lerman A (2003) Endothelial dysfunction: a marker of atherosclerotic risk. Arterioscler Thromb Vasc Biol 23:168–175 16. Giannotta M, Trani M, Dejana E (2013) VE-cadherin and endothelial adherens junctions: active guardians of vascular integrity. Dev Cell 26:441–454 17. Kavurma MM, Tan NY, Bennett MR (2008) Death receptors and their ligands in atherosclerosis. Arterioscler Thromb Vasc Biol 28:1694– 1702 18. Hansson GK, Chao S, Schwartz SM, Reidy MA (1985) Aortic endothelial cell death and replication in normal and lipopolysaccharidetreated rats. Am J Pathol 121:123–127 19. Stevens T, Garcia JG, Shasby DM, Bhattacharya J, Malik AB (2000) Mechanisms regulating endothelial cell barrier function. Am J Physiol Lung Cell Mol Physiol 279:L419– L422 20. Wojciak-Stothard B, Potempa S, Eichholtz T, Ridley AJ (2001) Rho and Rac but not Cdc42 regulate endothelial cell permeability. J Cell Sci 114:1343–1355 21. Hall A (1998) Rho GTPases and the actin cytoskeleton. Science 279:509–514 22. Machesky LM, Hall A (1996) Rho: a connection between membrane receptor signalling and the cytoskeleton. Trends Cell Biol 6:304–310

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23. Tzima E (2006) Role of small GTPases in endothelial cytoskeletal dynamics and the shear stress response. Circ Res 98:176–185 24. Braga VM, Machesky LM, Hall A, Hotchin NA (1997) The small GTPases rho and Rac are required for the establishment of cadherindependent cell-cell contacts. J Cell Biol 137: 1421–1431 25. Harrington EO, Shannon CJ, Morin N, Rowlett H, Murphy C, Lu Q (2005) PKCdelta regulates endothelial basal barrier function through modulation of RhoA GTPase activity. Exp Cell Res 308:407–421 26. Sanchez T, Skoura A, Wu MT, Casserly B, Harrington EO, Hla T (2007) Induction of vascular permeability by the sphingosine-1phosphate receptor-2 (S1P2R) and its downstream effectors ROCK and PTEN. Arterioscler Thromb Vasc Biol 27:1312–1318 27. Lu Q, Sakhatskyy P, Grinnell K, Newton J, Ortiz M, Wang Y, Sanchez-Esteban J, Harrington EO, Rounds S (2011) Cigarette smoke causes lung vascular barrier dysfunction via oxidative stress-mediated inhibition of RhoA and focal adhesion kinase. Am J Physiol Lung Cell Mol Physiol 301:L847–L857 28. Harrington EO, Brunelle JL, Shannon CJ, Kim ES, Mennella K, Rounds S (2003) Role of protein kinase C isoforms in rat epididymal microvascular endothelial barrier function. Am J Respir Cell Mol Biol 28:626–636 29. Lu Q, Harrington EO, Hai CM, Newton J, Garber M, Hirase T, Rounds S (2004) Isoprenylcysteine carboxyl methyltransferase modulates endothelial monolayer permeability: involvement of RhoA carboxyl methylation. Circ Res 94:306–315 30. Hamburger V, Hamilton HL (1992) A series of normal stages in the development of the chick embryo. 1951. Dev Dyn 195:231–272 31. Majumder S, Ilayaraja M, Seerapu HR, Sinha S, Siamwala JH, Chatterjee S (2010) Chick embryo partial ischemia model: a new approach to study ischemia ex vivo. PLoS One 5:e10524 32. Prozialeck WC, Edwards JR, Woods JM (2006) The vascular endothelium as a target of cadmium toxicity. Life Sci 79:1493–1506 33. Nagarajan S, Rajendran S, Saran U, Priya MK, Swaminathan A, Siamwala JH, Sinha S, Veeriah V, Sonar P, Jadhav V, Jaffar Ali BM, Chatterjee S (2013) Nitric oxide protects endothelium from cadmium mediated leakiness. Cell Biol Int 37:495–506 34. Saran U, Mani KP, Balaguru UM, Swaminathan A, Nagarajan S, Dharmarajan AM, Chatterjee S (2017) sFRP4 signalling of apoptosis and angiostasis uses nitric oxidecGMP-permeability axis of endothelium. Nitric Oxide 66:30–42

Chapter 16 Measurement of Transendothelial Electrical Resistance in Blood-Brain Barrier Endothelial Cells O’lisa Yaa Waithe, Xu Peng, Ed W. Childs, and Binu Tharakan Abstract The integrity of the blood-brain barrier (BBB), the protective barrier of the brain, is key to maintaining normal microvascular permeability and brain homeostasis. Brain microvascular endothelial cells are primary components of the blood-brain barrier. Transendothelial electrical resistance (TEER) is the electrical resistance across a cellular monolayer such as the brain microvascular endothelial cell monolayers. Measurement of TEER is considered a sensitive, reliable, and noninvasive method for evaluating barrier integrity and permeability of an endothelial cell monolayer under in vitro conditions. Measurement of TEER is also helpful for studying various cellular and molecular changes and signaling events that regulate barrier functions in endothelial monolayers. One of the in vitro endothelial cell barrier models that have been commonly used for measuring TEER is the BBB model using human or rodent brain microvascular endothelial cells grown as a monolayer. In this protocol, we describe how TEER is measured in brain microvascular endothelial cell monolayers, to determine blood-brain barrier integrity under in vitro conditions. Key words Transendothelial electrical resistance, TEER, Blood-brain barrier, Endothelial barrier permeability

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Introduction The endothelium, a monolayer of endothelial cells, constitutes the interior lining of the blood vessels and the lymphatic system. The endothelium plays a major role in physiological processes such as the control of vascular permeability, vasomotor tone, leukocyte trafficking between blood and underlying tissue, angiogenesis, and both innate and adaptive immunity [1]. Vascular endothelial cells provide a non-thrombogenic monolayer surface that lines the lumen of blood vessels, and its integrity is critical to normal vascular permeability and organ functions. In the blood-brain barrier (BBB) – the protective vascular barrier of the brain – the interendothelial tight junctions that consist of tight junction proteins regulate microvascular permeability. The BBB is a selectively permeable cellular barrier that separate apical (luminal) and basolateral

Binu Tharakan (ed.), Vascular Hyperpermeability: Methods and Protocols, Methods in Molecular Biology, vol. 2711, https://doi.org/10.1007/978-1-0716-3429-5_16, © Springer Science+Business Media, LLC, part of Springer Nature 2024

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(abluminal) sides in the brain, thereby controlling the transport processes to maintain brain homeostasis. Vascular barrier integrity is critical for the physiological activities of the tissue throughout the body [2–4]. In the body, vasoactive substances and mechanical stress triggered by hemodynamic forces, such as mechanical stretch and shear stress, stimulate endothelial cell signaling [5]. In the brain, dysfunction of the BBB results in vasogenic cerebral edema, causing elevated intracranial pressure (ICP), decreased cerebral perfusion, improper tissue oxygenation, brain herniation, and induction of apoptotic cell death [6–8]. Vascular endothelial barrier dysfunction at the level of the BBB occurs in various pathologies such as traumatic brain injury, and ischemic and hemorrhagic shock [7, 8]. Measurement of monolayer permeability and TEER are two of the reliable methods to determine barrier integrity and permeability to understand the abovementioned pathologies under in vitro conditions [9, 10]. Generally, transendothelial/epithelial electrical resistance reflects the ionic conductance of the paracellular pathway in the epithelial/endothelial monolayer, whereas monolayer permeability studies the flux of nonelectrolyte tracers and indicates the paracellular water flow, as well as the pore size of the tight or adherens junctions. In this chapter, TEER measurement from brain microvascular endothelial cell monolayers will be discussed [7, 8, 10–12]. The electrical resistance of an epithelial or endothelial monolayer, measured in ohms, is a quantitative measure of the barrier integrity and is an indirect indicator of permeability. The classical setup for measurement of TEER in endothelial monolayer, used in research settings, consists of a cellular monolayer cultured on a semipermeable filter insert that defines a partition for apical (or upper) and basolateral (or lower) compartments. For measuring the electrical resistance, two electrodes are used, with one electrode placed in the upper chamber and the second electrode in the lower chamber. The ohmic resistance is calculated based on Ohm’s law (i.e., current is directly proportional to voltage but inversely proportional to resistance) as the ratio of the voltage and current. The TEER measurement includes measuring the blank resistance (RBLANK) of the e-membrane only (without cells) and measuring the resistance across the endothelial cell layer on the semipermeable membrane (RTOTAL): RTOTAL includes the ohmic resistance of the endothelial cell layer RTEER, the endothelial cell culture medium RM, the semipermeable membrane insert RI, and the electrode medium interface REMI. The endothelial cell-specific resistance (RTISSUE), in units of Ω, can be obtained as follows: RTISSUE (Ω) = RTOTAL - RBLANK. The observed value is reported as follows: TEERREPORTED = RTISSUE (Ω) × Membrane (M)AREA (cm2). Resistance is inversely related to endothelial monolayer permeability.

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Materials 1. Human or rat brain (or any other microvascular endothelial cells) microvascular endothelial cells are suitable for this study. The method described in this protocol utilizes rat brain microvascular endothelial cells (RBMECs) and RBMEC medium. 2. Opti-MEM (MEM)/reduced serum medium and Dulbecco’s modified Eagle’s medium (DMEM; with high glucose, HEPES, without phenol red). 3. Phosphate-buffered saline (PBS). 4. Epithelial volt/Ohm Meter (EVOM2) together with STX2 electrode (World Precision Instruments). 5. Monolayer (Corning).

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Methods 1. Grow primary cultures of rat brain microvascular endothelial cells (RBMECs) on fibronectin-coated cell culture dishes, using rat brain endothelial media in optimal cell culture condition (95% O2, 5% CO2 at 37 °C) to 60–80% confluency. 2. Transfer the cells to Transwell inserts and grow for 72–96 h. 3. Treat the cells with the desired compound. 4. Place the apparatus on a heating pad at 37–40 °C. 5. Wash electrode and EndOhm chambers twice with deionized (DI) water. 6. Make sure all wires are connected. 7. Soak the upper and lower electrode in DMEM for 10 min (submerge top electrode with media). 8. Remove the media from the lower chamber of the EndOhm. 9. Add 1 mL of the desired media into the lower chamber of the EndOhm. 10. Remove the monolayer plate (containing the cells) from the incubator and place it next to EndOhm. 11. Take the cell monolayer inserts from the tray and place into the lower chamber of the EndOhm (please note that there needs to be enough space between the bottom of monolayer membrane and the bottom electrode), place the top electrode on the EndOhm, and ensure that it is submerged in the media of the monolayer chamber (see Notes 2 and 3). 12. Observe the level of the cell culture media in the lower chamber in relation to the level of media in the endothelial

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monolayer with the upper chamber with p electrode placed in it. Try to get the level of media in the lower chamber and in the monolayer chamber to somewhat equal levels (see Note 4). 13. When media levels are equal, wait for the readings on the EVOM2 to stabilize (same reading for 3–6 s) and then record the resistance value. 14. Calculate the final unit area resistance (Ω*cm2) by multiplying the sample resistance by the effective area of the membrane (see Note 5).

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Notes 1. The method described here describes the use of hydrogen peroxide as the test agent (Fig. 1). 2. There could be high readings by error if the top electrode is out of the media and in the air, or if there are bubbles on the top electrode (typically >12,000) on the EVOM2. 3. If there are any fluctuations in the readings, the chloride tips might need to be rechloritized. For this purpose, soak the electrodes in 5% bleach for 10–15 min (purple-black layer forms on electrodes).

Fig. 1 Transendothelial electrical resistance (TEER) measured in rat brain microvascular endothelial cell monolayers. Exposure of the monolayers to hydrogen peroxide (H2O2) resulted in significant concentration-dependent decrease in TEER values compared to untreated control monolayers ( p < 0.05; n = 6). (This figure is taken from our previously published article, Anasooya Shaji et al. [7]. Creative Commons License https://creativecommons. org/licenses/)

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4. For standardization of the procedures, typically 180 μL in upper chamber with 1 mL in lower chamber gives equal levels, but this may have to be modified based on different media and cell types used for the study. 5. Multiply by 0.33 cm2 for Costar Transwell inserts (or based on the insert specification).

Acknowledgments The authors acknowledge the support from the National Institutes of Health (NIH) grant 5 SC3 NS127765-02 (BT). References 1. Zielin´ska KA, Van Moortel L, Opdenakker G, De Bosscher K, Van den Steen PE (2016) Endothelial response to glucocorticoids in inflammatory diseases. Front Immunol 7:592 2. Sarah Y (2016) Signal transduction pathways in enhanced microvascular permeability. Microcirculation 6:395–403 3. Mehta D, Malik AB (2006) Signaling mechanisms regulating endothelial permeability. Physiol Rev 86:279–367 4. Sukriti S, Tauseef M, Yazbeck P, Mehta D (2014) Mechanisms regulating endothelial permeability. Pulm Circ4:535–551 5. Hirase T, Node K (2012) Endothelial dysfunction as a cellular mechanism for vascular failure. Am J Physiol Heart Circ Physiol 302:H499– H505 6. Logsdon AF, Lucke-Wold BP, Turner RC, Huber JD, Rosen CL, Simpkins JW (2015) Role of microvascular disruption in brain damage from traumatic brain injury. Compr Physiol 5:1147–1160 7. Anasooya Shaji C, Robinson BD, Yeager A, Beeram MR, Davis ML, Isbell CL, Huang JH, Tharakan B (2019) A tri-phasic role for hydrogen peroxide in blood-brain barrier endothelial cells. Sci Rep 9:133

8. Alluri H, Wiggins-Dohlvik K, Davis ML, Huang JH, Tharakan B (2015) Blood-brain barrier dysfunction following traumatic brain injury. Metab Brain Dis 30:1093–1104 9. Tinsley JH, Wu MH, Ma W, Taulman AC, Yuan SY (1999) Activated neutrophils induce hyperpermeability and phosphorylation of adherens junction proteins in coronary venular endothelial cells. J Biol Chem 274:24930– 24934 10. Robinson BD, Shaji CA, Lomas A, Tharakan B (2018) Measurement of microvascular endothelial barrier dysfunction and hyperpermeability in vitro. Methods Mol Biol 1717:237–242. https://doi.org/10.1007/978-1-4939-75266_19 11. 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 20:107–126 12. Vigh JP, Kincses A, Ozgu¨r B, Walter FR, SantaMaria AR, Valkai S, Vastag M, Neuhaus W, Brodin B, De´r A, Deli MA (2021) Transendothelial electrical resistance measurement across the blood–brain barrier: a critical review of methods. Micromachines (Basel) 12:685

Chapter 17 An In Vitro Bilayer Model of Human Primary Retinal Pigment Epithelial and Choroid Endothelial Cells for Permeability Studies Karthikka Palanisamy and Subbulakshmi Chidambaram Abstract The blood-retinal barrier (BRB) present in the posterior chamber of the eye plays a major role in maintaining the proper function and integrity of the retina. Retinal pigment epithelium and choriocapillaris form the outer blood retinal barrier (oBRB), and breakdown of this barrier leads to vision-threatening diseases like macular edema, macular degeneration, and diabetic retinopathy. A simplified cell culture model of oBRB will be of great importance in elucidating the molecular mechanism of the disease progression. This chapter describes methods for primary cell isolation from donor eyes to culture human retinal pigment epithelial cells (hRPE) and choroidal endothelial cells (hCEC) and the protocol for construction of a simplified in vitro model of oBRB on fibronectin-coated Transwell inserts. Further, we explained the permeability study using FITC-dextran conjugated tracers for validating the bilayer model. The permeability experiments ensured that the system could easily be manipulated to recapitulate the pathological condition in vitro. Thus, it would be an optimal system for studying the disease mechanisms related to retinal and choroidal pathologies, for screening small molecules, and for performing drug permeability kinetics. Moreover, fundamental understanding of paracellular and transcellular trafficking of cargo in hRPE and hCEC could also be studied using this model. Key words Donor eyes, Human primary choroidal endothelial cells, Human retinal pigment epithelial cells, In vitro blood retinal barrier model, Transwell inserts, FITC dextran permeability assay

1

Introduction The blood-retinal barrier (BRB) strictly regulates paracellular and transcellular transport of selected cargo by possessing specialized intercellular tight and adherent junction complexes. In particular, the retina must be able to regulate its osmotic balance, ion concentration, and nutrients while excluding immunoglobulins and immune cells. The retinal barrier is characterized by inner barrier (iBRB), comprised of retinal microvascular endothelial cells (hREC), and the outer barrier (oBRB) by retinal pigment epithelial (hRPE) cells located between the neural retina and Bruch’s

Binu Tharakan (ed.), Vascular Hyperpermeability: Methods and Protocols, Methods in Molecular Biology, vol. 2711, https://doi.org/10.1007/978-1-0716-3429-5_17, © Springer Science+Business Media, LLC, part of Springer Nature 2024

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membrane of the choroid. In diabetic retinopathy (DR), a number of studies have addressed the importance of iBRB, but the significance of oBRB has not been studied in detail. The effect of diabetes on oBRB had been emphasized, and the magnitude of RPE barrierspecific leakage in diabetic macular edema under retinopathy conditions had also been indicated [1, 2]. However, the lack of proper methodology to study RPE barrier in vivo is impeding the research on oBRB [2]. The cell culture models are essential tools in dissecting the molecular mechanism of disease pathology, but the selection of pertinent cell culture model is a critical issue [3]. Already existing in vitro models of oBRB could not entirely replicate the physiological biochemical milieu [4]. Many of the studies involving human oBRB either used ARPE-19 cell line or bovine RPE or porcine RPE along with human vein umbilical vein endothelial cells (HUVEC) or dermal endothelial or RF-6A cell lines, in the place of hRPE and hCEC, respectively [5–8]. In addition, there were striking differences among microvascular ocular endothelial cells. For instance, IGF-1 influenced proliferation of hREC but not hCEC [9]. The proteome of donor-matched retinal endothelial cells (hREC) and hCEC showed significant variations [10]. Hence, the species and tissue specificity of cells should be taken into account while deducing conclusions on the disease mechanism. In these lines, any functional assays using ARPE19 and HUVEC might not exactly typify the in vivo oBRB condition. In this chapter, we described the techniques involved in developing an in vitro outer blood-retinal barrier model consisting of hRPE, Transwell insert, and hCEC [11]. We optimized a simplified protocol for the isolation of primary hRPE and hCEC from donor eyes and characterized the cell types (Fig. 1). Then, we created an oBRB bilayer model using hRPE on basal side of the Transwell inserts and hCEC on the apical side (Fig. 2). Further, we validated the model using FITC dextran conjugated tracer permeability assay (Fig. 3). The overall workflow has been depicted in Fig. 4.

2

Materials

2.1 Cell Culture Media and Chemicals

1. Preparation of DMEM/F12 media for hRPE culture: Dulbecco’s modified eagle medium with Ham’s F12 nutrient mixture (DMEM (F12) medium (Cat no. 56498C, Sigma Aldrich) in powder form (without sodium bicarbonate) contains 3151 mg/L dextrose, 2.5 mM L-glutamine, with 55 mg/L of sodium pyruvate. It also contains inorganic salts, vitamins, amino acids, and other components necessary for cell growth. Dissolve DMEM/F12 powder in 900 mL of autoclaved sterile water, and add 2.44 g/L of sodium bicarbonate and mix thoroughly. Filter the solution using (Sartorius) membranefiltering unit having pore size of 0.2 μm (or less) under

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Fig. 1 Isolation and characterization of human primary ocular cells. (a) Top left panel shows human microvascular choroidal endothelial cells (hCEC) freshly isolated (P0) and the first passage (P1) on the right side. Bottom left and right panels show the human retinal pigment epithelial (hRPE) cells at P0 and P1, respectively. (b) Semiquantitative RT-PCR was done to show the presence of von Willebrand factor transcripts in hCEC and HUVEC. (c) RT-PCR for cytokeratin-18 in the primary hRPE and ARPE-19 cells was done. GAPDH was used as an internal control (n = 3). (d) Immunofluorescence staining of von Willebrand factor in hCEC and cytokeratin-18 in hRPE (n = 3). (e) Tube formation assay was done on Matrigel using hCEC. F. PEDF levels in the conditioned medium from hRPE grown insert (from upper and lower chamber) were measured by ELISA. P = 0.006 (n = 3). (g) VEGF ELISA was performed in the conditioned medium from the upper and lower chamber of hRPE, p = 0.02 (n = 3). (Reproduced from Ref. [11])

15–20 psi vacuum pressure. Aliquot the filtered medium in sterile 50 mL falcon tube under sterile conditions and store the media at 2–4 °C. Alternatively, DMEM/F12 media in liquid form can be used. 2. Preparation of endothelial growth media: EGM-2 basal medium (Cat no. CC3162, Lonza) along with bullet kit (Cat no. CC4176, Lonza) [containing FBS (10.00 mL), hydrocortisone (0.20 mL), hFGF-B (2.00 mL), VEGF (0.50 mL), R3-IGF-1 (0.50 mL), ascorbic acid (0.50 mL), hEGF (0.50 mL), GA-1000 (0.50 mL), heparin (0.50 mL)] was used for making endothelial growth media. The starvation media

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Fig. 2 Creating oBRB bilayer model of hCEC and hRPE on Transwell inserts. The hRPE cells were seeded onto the basal side of the insert kept upside down onto a 6-well plate. The cells were added with 100 μL of medium and the plate was kept half closed at 37 °C incubator with 5% CO2. After 4 h, the inserts were inverted and placed onto a 24-well plate containing epithelial medium. hCEC cells with endothelial medium were seeded on to the inner side of the inserts

contains EGM-2 basal medium with 1% FBS and no other supplements added. 3. EGM-MV (Cat no. 22020, PromoCell) containing fetal calf serum (0.05 mL/mL), endothelial cell growth medium (0.004 mL/mL), epidermal growth factor (10 ng/mL), ascorbic acid (90 μg/mL), hydrocortisone (1 μg/mL) was used for culturing hCEC cells to check the effect of VEGF. 4. Phosphate-buffered saline (PBS): To 800 mL of distilled water, add 8 g of NaCl, 0.2 g of KCl, 1.44 g of Na2HPO4, and 0.24 g of KH2PO4 and adjust the pH to 7.4 using HCl. Make up the volume to 1 L and sterilize PBS by autoclaving. 5. Heat-inactivate the fetal bovine serum (FBS) at 56 °C water bath for about 30 min. This ensures the inactivation of

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Fig. 3 Validation of the bilayer model. (a) Phase contrast image of cells in the bilayer culture and the inlet picture shows morphology of the primary hRPE and hCEC grown on inserts. (b) VEGF ELISA was performed from the conditioned medium taken from upper chamber of hRPE, hCEC, and hCEC/hRPE cultures (p = 0.02) (n = 3). (c) Permeability coefficient of 20 kDa FITC dextran in hRPE and hCEC monolayer treated with 100 ng/ml VEGF (n = 3) was measured. (d) The bilayer of hCEC/hRPE was treated with 100 ng/ml VEGF, and the permeability was calculated. After 2 h, anti-VEGF agent bevacizumab (0.125 mg/ml) was added and the permeability was measured for further 2 h (n = 3). (Reproduced from Ref. [11])

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Fig. 4 Schematic diagram of the bilayer model and permeability assay. The hRPE and hCEC cells were isolated from donor eyes. The primary cells were grown on either side of the Transwell insert. To create the pathological milieu, these bilayer cells were treated with VEGF and their barrier permeability was checked using dextran-conjugated FITC flux. (Reproduced from Ref. [11])

complement proteins (involved in immune response). Swirl the bottle once after 15 min. 6. Antibiotic mix: Antimycotic solution (100×) (Cat no. 15240062, Gibco life technologies) containing 10,000 units/mL of penicillin, 10,000 μg/mL of streptomycin, and 25 μg/mL of Gibco Amphotericin B was used in 1× concentration. 7. Collagenase type II. 8. Fibronectin (Cat no. F2006, Sigma Aldrich). 9. 1% gelatin (w/v) prepared from adding 1 g of gelatin to 100 mL of PBS, after mixing properly, it should be autoclaved and stored at 4–8 °C. 10. Bovine serum albumin (BSA) for cell culture. 11. Tracers: FITC dextran 20 kDa, 1.2 mg/ml of final concentration was used for the experiments. 12. Anti-CD-31 coated Dynabeads (Cat no. 11155D, Invitrogen) with magnetic separator. 13. In vitro angiogenesis assay kit containing Matrigel and buffer.

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14. 0.1%, 0.25%, and 1.5% trypsin-EDTA (0.1 g or 0.25 g or 1.5 g of trypsin, 0.5 g of Dextrose and 0.03 g of EDTA [0.1 mM EDTA]) solution prepared in PBS (autoclaved) and filter sterilized before use. 15. Anti-VEGF (bevacizumab). 2.1.1 Cell Culture Plasticware and Other Accessories

1. 24-well plates (Corning), 6.5 mm polyester membrane Transwell insert with 0.4 μm pore size (Corning, CLS3470); Falcon tubes; 15 mL, 50 mL, and 40 mm culture dishes; 96-well plates; 25 cm2 and 75 cm2tissue culture flasks; and cryopreservation tubes. 2. 21G sterile needle, glass tissue homogenizer, sterile scalpel, autoclaved scissors, and forceps.

2.2

ELISA Kits

1. PEDF and VEGF ELISA kits (DY1177-05 and DVE00, R&D systems). 2. Multichannel pipette, wash buffer plate, and fresh/sterile microtips.

2.3 Immunofluorescence Antibodies, Chemicals, and Plastics

1. 24-well plates, 12 mm diameter coverslips (sterilized with ethanol before use). 2. TritonX-100 (SRL, cat no. 20202130), paraformaldehyde (prepare 4% and store at 4 °C), BSA and PBS for blocking solution, DAPI (5 mg/mL) and 50% glycerol. 3. Antibodies: rabbit raised-Anti-vWF (1:400); (cat no. A0082, Dako), mouse raised-anti-cytokeratin-18 (1:200); (cat no. 4548, cell signaling technologies (CST)). 4. Secondary anti-rabbit (Cat no. 4412, CST) and anti-mouse Alexa 488 (Cat no. 4408, CST) antibodies. 5. Images were taken in Carl Zeiss pseudoconfocal microscope.

2.4

RT PCR Materials

1. For RNA isolation: chloroform and isopropanol, ethanol. 2. 0.1% of diethyl pyrocarbonate (DEPC)-treated water. 3. iScript cDNA synthesis kit (Cat no. 1708891, Biorad). DNTP’s 10 nM, 10× Taq buffer with 20 mM MgCl2, Taq polymerase (Sigma Aldrich), DNA molecular weight marker (100–600 kb) and Milli-Q water. 4. 1.5 mL microcentrifuge tips, 0.2 mL PCR tubes. 5. 10 × TBE: Dissolve Tris (108 g) and boric acid (55 g) in 900 mL of water; prepare 0.5 M Na2EDTA and add 40 mL to the above solution and adjust the pH to 8, make up the total volume to 1000 mL and store at room temperature. Prepare 1× TBE for casting and running agarose gel.

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6. Agarose (electrophoresis grade, Sigma Aldrich), prepare stock solution of 10 mg/mL ethidium bromide (Sigma Aldrich) solution and add final concentration of 0.5 μg/mL to the agarose gel.

3

Methods

3.1 Coating the Culture Dishes with Extracellular Matrix Material

1. Coat the culture dishes using 1 mL of fibronectin solution (10 μg/mL dissolved in 1× PBS). 2. Incubate the coated dishes at 37 °C with 5% CO2 incubator for 20 min. 3. Remove the fibronectin solution and air dry the culture dishes under sterile conditions.

3.2 Preparation of Cadaver Eye Tissues Prior to Cell Isolation

1. The eyecups were enucleated within 6 h of death and processing should be done within 2 h of procurement for best results. 2. A total of three eyecups were used in this study (32- and 35-year-old male and 42-year-old female donors) (see Note 1). 3. Wash the procured eyecups twice with PBS added with 1× antibiotic-antimycotic solution (stabilized with 10,000 units of penicillin, 10 mg streptomycin and 25 μg amphotericin B per mL) and incubate in fresh solution for about 15 min.

3.2.1 Isolation and Culturing of Retinal Pigment Epithelial Cells from Human Eyecups

1. The eyecups were taken out from the PBS-antibiotic solution and placed on a sterile Petri plate. The isolation procedure was conducted with few modifications from Zhu et al. [2]. 2. Make a careful incision on the corneal layer near iris portion and continue the incision to make a circle cut. Open the eyecups such that the iris and ciliary portion will be clearly visible for its removal. 3. After complete removal of the iris and ciliary portion, remove the vitreous gel using two forceps (like pulling out a rope). 4. Once confirming the removal of vitreous gel completely, separate the retina carefully without disturbing the RPE layer (see Note 2). 5. Now cut the eyecup at four sites (making a cross-cut). This cut should be made deep such that the cut reaches till the optic nerve head. 6. Once the cut is made, spread the pieces like a floral section. Now carefully cut the single floret and place it on the fibronectin-coated culture dishes such that the RPE side is facing the bottom. 7. Now slowly remove the sclera holding the lower RPE layer with forceps (see Note 3). Leave the explants undisturbed for

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about 4–5 h and slowly add 100 μL of EGM-2 media above the explant to prevent it from drying. 8. Incubate the culture dishes with explants at 37 °C with 5% CO2 incubator overnight. On the next day, remove the explants slowly (see Note 4) and add 3 mL EGM-2 media (see Note 5). 9. Change the medium on alternate days until the growth of RPE is visible from the remnants of the explants attached to the plate. Continue changing the medium on alternate days until the culture becomes confluent (not monolayer). 10. Trypsinize the RPE cells using 0.25% trypsin solution and transfer to 0.1% sterile filtered gelatin-coated 25 cm2tissue culture flasks for further growth. Now change the medium from EGM-2 to DMEM-F12 with 10% FBS (from passage 1 (P1)). 11. For later passages use 0.1% trypsin-EDTA solution to trypsinize the cells. 12. In passage 2 (P2), seed the cells into 2 or 3 (75 cm2) tissue culture flasks according to the confluence of the culture in P1 (400,000 cells/75 cm2 flask). 13. After P2, the cells can be used for characterization studies and other experiments or can be cryopreserved for further use. 14. Use up to passage 5 for the experiments. 3.2.2 Isolation of Choroid Microvascular Endothelial Cells from Human Eyecups

1. The isolation of choroid microvascular endothelial cells starts with steps similar to isolation of hRPE (Subheading 3.2.1) until step 4. After that remove the RPE/BM/choroid layer gently from the eyecup. 2. Keep RPE/BM/choroid layer in a new sterile Petri plate such that RPE is upside down and wash it with PBS (see Note 6). 3. Now mince the washed RPE/BM/choroid layer into small pieces and add 1 mL of PBS (see Note 7) and transfer the minced pieces with PBS to a fresh 15 mL falcon tube. 4. Centrifuge the minced RPE/BM/choroid at 240 g for 3 min. This step will remove PBS and the tissue will be ready for trypsin treatment. 5. Further, add 2 mL of 1.5% trypsin-EDTA solution to minced pieces and incubate for 20 min at 37 °C water bath. 6. After trypsin-EDTA treatment, homogenize the minced RPE/BM/choroid pieces in a glass homogenizer and aspirate through 21G syringe needle for complete disassociation of the cells (see Note 8). 7. Now treat the homogenate with 1 mL of 0.2 mg/mL of collagenase type II for about 40 min at 37 °C water bath.

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8. Stop the collagenase action with DMEM F12 with 10% FBS (see Note 9). 9. Later incubate with anti-CD31-coated Dynabeads for about 1 h at room temperature in a rocker kept under sterile conditions. 10. Using a magnetic separator, isolate the CD-31-positive endothelial cells. 11. Suspend the endothelial cells along with beads in 2 mL of EGM-2 media and then seed on to fibronectin-coated 6 mm culture dishes. 12. Give media change daily (slowly over the sides of the dish), and after 2 days, visible growth of choroid endothelial cells can be seen under microscope (see Note 10). 13. Once the growth of endothelial cells reaches confluence (not monolayer), trypsinize and seed the cells onto 0.1% sterile gelatin-coated 25 cm2 flasks. 14. Similar to hRPE, use hCEC passages between P2 and P5 for further experiments and the rest can be cryopreserved. 3.3 Characterization of Isolated Primary Culture 3.3.1

RNA Isolation

1. Wash the cells grown on culture dishes or 25 cm2tissue culture flasks once with PBS, and treat with 1 mL or 2 mL of TRI reagent respectively for about 2 min and mix by repeated aspiration. 2. Transfer the TRI reagent to sterile DEPC-treated 1.5 mL vials and proceed with RNA isolation. 3. Add 200 μL of chloroform to 1 mL of TRI reagent with cells and shake well and let it stand for 15 min at room temperature. Now centrifuge at 12,000g for 15 min at 4 °C to separate the aqueous layer containing the RNA into new 1.5 mL centrifuge tubes. 4. Add 500 μL of isopropanol to the aqueous layer to precipitate RNA, gently mix the tube, and let it stand for 5 min and centrifuge at 12,000g for 10 min at 4 °C. The RNA pellet will be formed at the bottom of the vial. 5. Carefully remove the supernatant and wash the RNA pellet with 1 mL of 70% ethanol solution prepared with sterile DEPC-treated water. Let the tube stand for 5 min at room temperature and centrifuge at 14,000g for 5 min at 4 °C. 6. Discard the ethanol and keep the tubes to air dry. 7. Dissolve the pellets with 20 μL DEPC water and quantify using nanodrop or spectrophotometer and RNA should have A260/ A280 ≥ 1.7 (which indicates its purity without DNA and protein contamination).

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1. Take 100 ng of RNA and convert it into cDNA using i-script cDNA synthesis kit using manufacturer’s protocol. 2. Briefly, take 100 ng of RNA in PCR tubes along with 4 μL of iScript mix containing oligo dT and random primers along with 1 μL of reverse transcriptase enzyme. 3. Adjust the total reaction volume to 20 μL using nuclease-free water provided in the kit and set the reaction for 40 min in a PCR machine. 4. The program for cDNA conversion is as follows:

5. After cDNA conversion, set the PCR reaction using respective primers for housekeeping and the target genes along with negative control (no template). 6. Select the PCR primers 100% complementary to cytokeratin-18 (NM_199187.1) and vWF (NM_000552.3). The primer sequence of cytokeratin-18; FP: 5′- CACACAGTCTGCT GAGGTTG -3′, RP: 5′- TAAAGTCCTCGCCATCTTCC -3′ and vWF; FP: 5′- ACATCTTCACATTCACTCCACAAAA3′, RP: 5′-ATTCCTGAACAAGTGTTTTCCAGTC-3′. 7. Set the PCR reaction for a total volume of 25 μL, which includes 4 μL DNTP, 2.5 μL 10× buffer, 1 μL each FP and RP (100 picomole concentration), 0.3 μL Taq polymerase, 14.2 μL MilliQ water, and 2 μL cDNA.

3.3.3 Gel Run and Imaging

8. Since the annealing temperature for both cytokeratin-18 and vWF was 58.3 °C, use the same PCR program for both as given below. 1. Prepare 1.8% agarose gel in 1× TBE solution and microwave the solution until the agarose dissolves. 2. Add few drops of ethidium bromide (see Note 12) once the agarose mixture comes to 60–70 °C and pour into gel loading

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tray. Choose the tray and comb according to the sample quantity. 3. Place the comb before the gel gets solidified or the comb can be placed before pouring the gel too. 4. Once the gel gets solidified, place the gel tray in 1XTBE-filled buffer tank after removing the comb carefully. Add 5× loading dye to the PCR product and mix well before loading. Load the molecular weight ladder and negative control adjacent to the samples. Positive controls taken for this experiment include HUVEC DNA for vWF and ARPE-19 DNA for cytokeratin-18. 5. Run the gel at 50 V and capture the images using FluorChem FC3 (ProteinSimple). 6. Result: Agarose gel image showed the presence of vWF in hCEC and cytokeratin-18 in hRPE (Fig. 1b, c). 3.3.4 Immunofluorescence Imaging

1. Culture the isolated primary ocular cells (35,000 cells) over coverslips placed on 24-well plate. 2. After 11 days of growth, the cells will be ready for immunofluorescence study. 3. Remove the culture media and wash the cells once with PBS. 4. Fix the cells with 4% paraformaldehyde for about 7 min at room temperature. 5. Aspirate the paraformaldehyde and wash the cells three times with PBS. 6. Permeabilize the cells with 0.1% Triton X-100 in PBS for about 5 min at room temperature. 7. Block the cells with 3% BSA in PBS for 1 h at room temperature in the rocker. 8. Prepare primary antibody according to manufacture protocol in blocking solution and add to the cells. Incubate overnight at 4 °C. 9. On the next day, wash the cells, add secondary antibody, and incubate for 2 h at room temperature. 10. Wash the cells three times for 5 min each. 11. Add DAPI for staining the nuclei. Incubate the cells with DAPI for about 5 min at room temperature in the rocker. 12. Now wash the coverslips three times for 5 min each. 13. Prepare 50% glycerol with PBS and add a drop to the sterile slides, and mount the coverslip carefully upside down using forceps (see Note 13). 14. The images were taken using Carl Zeiss pseudo confocal microscope.

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15. Result: The fluorescence staining confirmed the purity of the culture at every passage along with phase contrast image showing the cobble stone appearance of hCEC and pigmented polygonal honeycomb appearance of hRPE (Fig. 1a). In the immunofluorescence study, hCEC showed positive staining for endothelial specific marker vWF, and hRPE showed positive staining toward cytokeratin-18 (Fig. 1d). 3.3.5 Tube Formation Assay

1. On the prior day of experiment, keep the Matrigel and buffer vial (-20 °C storage) at 4 °C for slow thawing (without affecting the growth factors present in it). 2. Keep a 96-well plate and 200 μL cut tips (edges are cut for about 1 cm using sterile scalpel) in sterile tip box in the freezer for about an hour. 3. During the experiment, mix the required buffer to Matrigel solution and place the vial in ice. This procedure is done in a sterile environment. 4. Place the 96-well plate on ice and add about 45 μL of Matrigel with buffer using the cooled cut tips and make sure it doesn’t have bubbles; if bubble appears, strike it with a sterile needle (see Note 14). 5. Now keep the plate inside 37 °C incubator with 5% CO2 for about 1 h. 6. Meantime, trypsinize hCEC cells grown in 25 cm2 flask using 0.1% trypsin: EDTA solution and centrifuge at 1500g for 3 min. Remove trypsin solution and add fresh EGM medium and count the cells. 7. Take 15,000 hCEC cells in 50 μL medium and add 150 μL of EGM-2 media (making the total volume to 200 μL in each well). 8. Add the medium with cells to Matrigel poured wells. Always check before adding the cells if the Matrigel is completely set. 9. A positive control should be added with 10 ng of VEGF/ml. 10. Incubate the plates at 37 °C incubator with 5% CO2 for about 4 h. 11. Check for the development of tubes in positive control wells, and take the pictures of hCEC tubes under phase contrast microscope. 12. Result: As a part of the functional characterization, the isolated hCEC cells formed distinct tubes (Fig. 1e).

3.4 Bilayer Culture on Transwell Inserts

1. Take the sterile Transwell inserts (24-well plate 0.4 μm pore size, polycarbonate) for bilayer culture. The inserts are placed upside down (inverted position) in a 6-well plate under sterile conditions (Fig. 2).

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2. Coat the bottom side of the insert with 0.1% gelatin and keep at 37 °C incubator with 5% CO2 for about 20 min. 3. Remove the gelatin coating from the insert and seed hRPE (106 cells /ml) with 100 μL of DMEM F12/10% FBS medium. Keep the lid covered halfway through. The seeded cells are left undisturbed and kept in 37 °C incubator with 5% CO2 for about 4–6 h. 4. Then, flip the inserts onto a 24-well plate filled with DMEM/ F12 with 10% FBS medium. 5. Remove the medium from the bottom and add 500 μL of fresh DMEM F12 with 10% FBS medium. This step will remove the unattached hRPE cells from the basal part of the insert. 6. Now the choroidal endothelial cells (106 cells/ml) can be seeded onto the inner part of the inserts (apical side) with 200 μL of EGM-MV (see Note 15). 7. Allow the cells to grow for the next 11 days until further experiment, and replace media every 2 days (with respective medium for hRPE and hCEC). 3.5 ELISA for PEDF and VEGF

1. Take the conditioned medium from hRPE, hCEC, and hCEC/ hRPE (see Subheading 3.4) grown on apical and basal sides of Transwell inserts. 2. Centrifuge the medium for about 1500g for 5 min and then store the conditioned medium at -80 °C until use. This centrifugation will remove the dead cells from the medium. 3. Keep the Elisa kits for PEDF and VEGF out for 30 min prior to the experiment. 4. Follow the manufacturer’s protocol. The standards, reagents, and samples were prepared according to the protocol and take required amount of microplate strips and store the remaining immediately at 4 °C. 5. Add 50 μL assay diluent to each well along with conditioned media and controls. Add 200 μL standards and incubate for 2 h at room temperature. 6. Record the order of samples in the layout given by the provider to avoid mistakes. 7. Aspirate the solution and wash the wells with 400 μL of 1× wash buffer prepared from 25× stock and wash using multichannel pipette. 8. Wash three times and aspirate the wash buffer and tap the plate upside down in a dry tissue. 9. Now add 200 μL VEGF conjugate and incubate for 2 h at room temperature. 10. Repeat the wash as above for three times.

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11. Prepare substrate solution and add 200 μL in each well and leave in dark covered for about 20 min at room temperature. 12. Add 50 μL of stop solution and ensure thorough mixing for proper reading. 13. Measure the optical density at 450 nm and subtract from 540 nm or 570 nm to correct the optical imperfection from the plate. 14. The readings are used to calculate the value of VEGF and PEDF from standards and represented in pg/ml and ng/ml respectively. 15. Result: PEDF level was high on the apical side of the inserts, whereas VEGF level was significantly high on the basal side of the hRPE grown insert (Fig. 1f). This specific secretory property confirms the polarity of hRPE grown on the inserts [12]. Further, the levels of VEGF were found to be increased from hCEC grown in bilayer than grown as monolayer on the inserts. This shows that bilayer was able to mimic the outer barrier system with proper secretory function (Fig. 3b). 3.6 Paracellular Permeability Assay Using the Bilayer Model

1. To check the effect of VEGF, grow hRPE with DMEM with 10% FBS and hCEC in EGM-MV medium without VEGF, as monolayer and bilayer cultures. 2. Give 3 h of serum starvation for monolayer and bilayer cultures with DMEM 1% FBS and EBM 1% FBS for hRPE and hCEC, respectively. 3. Add 100 ng/ml of VEGF to the cultures and treat for 3 h at 37 °C incubator with 5% CO2. 4. Add 20 kDa FITC dextran to the upper chamber (inner part) of the insert at the concentration of 1.25 mg/mL (in 50 μL) (see Note 15). 5. Before taking basal reading, keep the Transwell insert plates at room temperature under sterile condition for about 20 min. 6. Take an amber 96-well plate for measuring the fluorescence readings. Add 100 μL of serum starvation medium as blank. Repeat this for every reading. 7. Now take 100 μL of medium from the lower chamber of the insert by slowly lifting the insert upward using a sterile forceps. Ensure proper mixing of the fluorescence tracers in the medium by repeated slow pipetting before taking the sample out for fluorescence reading. 8. Measure the fluorescence using SpectraMax M2e with excitation at 485 nm and emission at 535 nm. 9. Replace the medium to the respective wells after taking the reading and keep the plates in 37 °C incubator with 5% CO2 for

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1 h, and reading should be taken every hour for about 4 h repeating the same steps with separate blank each time. 10. With respect to bilayer, at the end of 2 h, add anti-VEGF agent (bevacizumab) to the upper chamber of the inserts to check VEGF counter effect. 11. Take the readings after 1 h of incubation at 37 °C incubator with 5% CO2. 12. Repeat the same procedure for the second hour reading. 13. Calculate the apparent permeability coefficient (Papp) using the following formula: P app =

dQ 1 ðcm=sÞ dt A
C 0
60

where dQ/dt is the amount of FITC transported per minute (ng/min), A is the surface area of the filter (cm2), C is the initial concentration of FITC (ng/ml), and 60 is the conversion from minutes to seconds [13]. 14. Result: Both hRPE and hCEC monolayers showed increased permeability with 100 ng/mL of VEGF. The bilayer culture showed statistically significant increase in the permeability with VEGF [14]. Further, with anti-VEGF added to the inserts counteracted the VEGF-induced permeability significantly (Fig. 3d).

4

Notes 1. The success of hRPE and hCEC isolation largely depends on the age of donor eyecups and processing speed. The younger the eyecup, the higher the yield will be. 2. Make sure there are no remnant pieces of retina lying above the RPE layer since other retinal cells might grow by attaching the culture dish, since we use endothelial-specific medium initially for faster growth. Thus, it is important to make sure there is no retina attached. 3. Since this step is done after the RPE layer settles on the culture dish, removal of sclera should be done very carefully as abrasive removal might damage the cells at the bottom or may collapse the spreading of RPE layer. 4. While removing the explants, gently pull upward such that it doesn’t disturb the attached RPE cells. 5. Two things have to be noted while giving change of medium for hRPE: (1) give media change over the edges of the plate since the attached cell from the explants will get disturbed if change of medium is given hard and fast. (2) Check for the

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presence of any contamination. Even after washing the cadaver tissues in antibiotic-antimycotic solution, there are chances of bacterial growth in the explant method. If present, give a wash using PBS with antibiotic-antimycotic solution followed by two washes with growth medium with 1× antibioticantimycotic solution, and then add fresh EGM-2 medium. Check after 4–6 h of the washing step, and if the bacterial growth does not subside, discard the culture and start fresh since this step is important to get a contamination-free culture for further experiments. 6. During washing step in hCEC isolation, the RPE layer should be at the top and this will remove RPE cells to some extent. 7. The RPE/BM/choroid tissue can be minced in the Petri plate, or the tissue with PBS can be transferred to a sterile 50 mL falcon tube and minced with surgical scissors. Adding PBS will help in cutting the pieces with ease. 8. Collagenase treatment and passing through syringe needle will aid in releasing the endothelial cells out of the microvascular blood vessels. So make sure you do these steps with utmost patience and perfection, and this will help the success of the cell isolation. 9. The addition of DMEM/F12 with 10% FBS is just to stop the collagenase action. This step can be done with addition of FBS alone as well. 10. The growing choroidal endothelial cells will initially be seen along with the beads, but this is not to be worried as the beads will be removed in the subsequent trypsinization steps and further passages will be free of magnetic beads. 11. Since ethidium bromide is a potent carcinogenic, be careful while adding it to the agarose gel. If you add ethidium bromide when the agarose mixture is hot, the chances are high that you inhale the fumes. 12. Before mounting the coverslips on the slides, just tap them by holding at a 45° angle with a dry tissue. This will remove any liquid present on the coverslips which will facilitate proper mounting. 13. The Matrigel should be released slowly by touching the bottom of the plate to avoid bubbles. If bubbles appear, it should be poked with a sterile needle. Don’t use the same needle for two bubbles since by capillary action, gel mixture will enter the needle and poking the next bubble will be difficult and meanwhile the gel may set. 14. For checking the effect of VEGF, hCEC should be grown with EGM-MV media, which does not have VEGF, from the day it has been seeded onto the inserts.

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15. Always prepare fresh FITC solution and do not prepare two sets of FITC solution for different wells in one experiment. Moreover, prepare FITC solution in high concentration (say >5 mg) so that it will be easy for proper mixing of the FITC in the medium used.

Acknowledgments We would like to acknowledge the financial assistance provided by Department of Biotechnology under the project “DBT-RGYI BT/PR15023/GBD/27/282/2010,” DBT-Neurobiology Taskforce grant BT/PR15023/GBD/27/282/2010, and Max Planck-India mobility grant “IGSTC/MPG/FS (SC) 2012.” CS thanks the support from University Grants Commission, New Delhi, for the award of Assistant Professorship under its Faculty Recharge Program (UGC-FRP). References 1. Simo´ R, Villarroel M, Corraliza L, et al (2010) The retinal pigment epithelium: something more than a constituent of the blood-retinal barrier-implications for the pathogenesis of diabetic retinopathy. J Biomed Biotechnol 1–15. https://doi.org/10.1155/2010/ 190724 2. Zhu M, Med M, Provis J, Penfold P (1998) Isolation, culture and characteristics of human foetal and adult retinal pigment epithelium. Aust New Zeal J Opthalmol 26(Suppl 1): S50–S52. Epub 1998/07/31 3. Kuznetsova A V., Kurinov AM, Aleksandrova MA (2014) Cell models to study regulation of cell transformation in pathologies of retinal pigment epithelium. J Ophthalmol 1–18. https://doi.org/10.1155/2014/801787 4. Cai H, Del Priore LV (2006) Gene expression profile of cultured adult compared to immortalized human RPE. Mol Vis 12:1–14 5. Mannermaa E, Reinisalo M, Ranta V-P et al (2010) Filter-cultured ARPE-19 cells as outer blood-retinal barrier model. Eur J Pharm Sci 40:289–296. https://doi.org/10.1016/j.ejps. 2010.04.001 6. Chung M, Lee S, Lee BJ et al (2018) Wet-AMD on a chip: modeling outer bloodretinal barrier in vitro. Adv Healthc Mater 7:1– 7 . h t t p s : // d o i . o r g / 1 0 . 1 0 0 2 / a d h m . 201700028

7. Maugeri G, D’Amico AG, Rasa` DM et al (2017) Caffeine prevents blood retinal barrier damage in a model, in vitro, of diabetic macular edema. J Cell Biochem 118:2371–2379. https://doi.org/10.1002/jcb.25899 8. Spencer C, Abend S, McHugh KJ, SaintGeniez M (2017) Identification of a synergistic interaction between endothelial cells and retinal pigment epithelium. J Cell Mol Med 21: 2542–2552. https://doi.org/10.1111/jcmm. 13175 9. Browning AC, Halligan EP, Stewart EA et al (2012) Comparative gene expression profiling of human umbilical vein endothelial cells and ocular vascular endothelial cells. Br J Ophthalmol 96:128–132. https://doi.org/10.1136/ bjophthalmol-2011-300572 10. Smith JR, Choi D, Chipps TJ et al (2007) Unique gene expression profiles of donormatched human retinal and choroidal vascular endothelial cells. Investig Ophthalmol Vis Sci 48:2676–2684. https://doi.org/10.1167/ iovs.06-0598 11. Palanisamy K, Karunakaran C, Raman R, Chidambaram S (2019) Optimization of an in vitro bilayer model for studying the functional interplay between human primary retinal pigment epithelial and choroidal endothelial cells isolated from donor eyes. BMC Res Notes 12. https://doi.org/10.1186/s13104019-4333-x

An in vitro Bilayer Model of Human Blood-Retinal Barrier 12. Sonoda S, Sreekumar PG, Kase S et al (2010) Attainment of polarity promotes growth factor secretion by retinal pigment epithelial cells: relevance to age-related macular degeneration. Aging (Albany NY) 2:28–42 13. Ma X, Zhang H, Pan Q et al (2013) Hypoxia/ aglycemia-induced endothelial barrier dysfunction and tight junction protein downregulation can be ameliorated by citicoline. PLoS One 8:

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4–9. https://doi.org/10.1371/journal.pone. 0082604 14. Klaassen I, Van Noorden CJF, Schlingemann RO (2013) Molecular basis of the inner blood e retinal barrier and its breakdown in diabetic macular edema and other pathological conditions. Prog Retin Eye Res. https://doi.org/10. 1016/j.preteyeres.2013.02.001

Chapter 18 Quantifying Adhesion of Inflammatory Cells to the Endothelium In Vitro Saravanakumar Muthusamy Abstract We present a simple and quantitative assay system that accurately models human endothelium by use of primary human umbilical vein endothelial cells (HUVECs) in cell culture plates coated with gelatin, a matrix that mimics basal lamina, the matrix that is tightly associated with the vascular endothelium and is critical for its proper function. We describe using this system to quantitatively measure adhesion of the inflammatory cells – monocytic THP-1 cell line to the HUVEC monolayer. The THP-1 cells are fluorescently labeled which allows to quantify the number of the fluorescent THP-1 cells adhering to the endothelium under a microscope and the level of florescence – a quantitative measure of the number of adhering fluorescent THP-1 cells using a fluorescent plate reader. After optimization, we were able to detect increased adhesion of the THP-1 cells to the endothelium in response to the inflammatory cytokine TNFα in a dose-dependent manner like what has been observed in vivo. Key words Endothelium, Leucocytes, Adhesion, Inflammation, Extravasation, Tumor metastasis, Tumor inflammation

1

Introduction Extravasation of inflammatory cells out of the circulatory system through the endothelial monolayer of the vasculature toward the site of tissue damage in inflammation or site of infection is a major early step in inflammatory responses in atherosclerosis [1], sepsis, and tumor inflammation [2]. Similarly, extravasation of tumor cells to sites of secondary metastasis is a major step in tumor metastasis [3, 4]. The first step in the process of extravasation is the adhesion of the inflammatory cells [1, 2] or tumor cells [5] to the endothelial monolayer as a result of endothelial activation. The adhesion is mediated by activation-induced interactions between endothelial cell adhesion molecules and their specific ligands on the inflammatory cells [6] or the tumor cells [7]. Therapeutic molecules that inhibit the process of adhesion of inflammatory cells to the endothelium are well-known anti-

Binu Tharakan (ed.), Vascular Hyperpermeability: Methods and Protocols, Methods in Molecular Biology, vol. 2711, https://doi.org/10.1007/978-1-0716-3429-5_18, © Springer Science+Business Media, LLC, part of Springer Nature 2024

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inflammatory drugs [8] and inhibiting the interaction of tumor cells with endothelial cells has been shown to prevent metastasis [7]. However, many of the anti-adhesion therapy clinical trials for inflammation have been disappointing [9] and antimetastatic therapies targeting inhibition of adhesion of tumor cells to the endothelium are lacking [10]. To develop more effective anti-adhesion therapies for inflammation and cancer, a better understanding of the molecular mechanisms controlling the adhesion process is needed which requires to have a quantitative adhesion assay in a system that mimics adhesion of cells to the human endothelium in vivo. We present an assay system that accurately models human endothelium by use of primary human umbilical vein endothelial cells (HUVECs) in cell culture plates coated with gelatin, a matrix that mimics basal lamina that is tightly associated with the vascular endothelium and is critical for its proper function [11]. We describe using this system to quantitatively measure adhesion of the inflammatory cells – THP-1 cells to the HUVEC monolayer. After optimization, we were able to detect increased adhesion of the THP-1 cells to the endothelium in response to the inflammatory cytokine TNFα in a dose-dependent manner similar to what has been observed in vivo. We have thus confirmed the relevance of this assay to quantitatively measure adhesion of inflammatory cells to the endothelium. Further, this assay can be easily adapted to quantify adhesion of tumor cells to the endothelium, which would help defining the molecular mechanisms underlying tumor cell adhesion and metastasis and testing potential drugs that inhibit tumor cell adhesion as antimetastatic therapies. A schematic diagram of the assay with various steps of the procedure is included (Fig. 1).

2

Materials 1. 48-well tissue culture plates. 2. Gelatin solution (0.1% in PBS). 3. PBS. 4. Endothelial cells (HUVEC – human umbilical vein endothelial cells). 5. EGM-2 Media (LONZA, Catalog # CC-3162). 6. EBM- 2 media (LONZA., Catalog # CC-3156). 7. Fetal bovine serum (FBS). 8. TNFα. 9. Leukocytes (THP1 – monocytes). 10. RPMI1640 media.

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Endothelial monolayer on gelan coated plate

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Leukocytes grown in suspension Fluoroscent labelling of Leukocytes

Endothelial acvaon (TNFα)

Pouring of labeled Leukocytes to the endothelial monolayer + 1 hr Incubaon

Washing out non adhering cells

Lysis of Leukocytes to release labeling dye

Counng of adhered Leukocytes under microscope

Quanficaon of Fluorescent dye on plate reader

Fig. 1 The adhesion assay principle

11. 500× green, fluorescent labeling dye (Cell Biolabs Inc., Part No. 12101). 12. 4× cell washing buffer (Cell Biolabs Inc., Part No. 10404).

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13. 10× cell lysis buffer (Cell Biolabs Inc., Part No. 12104). 14. BSA. 15. Cell culture incubator (37 °C, 5% CO2 atmosphere). 16. Light microscope. 17. 96-well plates suitable for a fluorescence plate reader. 18. Fluorescence plate reader.

3

Methods

3.1 Gelatin Coating of 48-Well HUVEC Culture Plates

1. Under sterile conditions, add 200 μL of 0.1% gelatin solution to each well of a 48-well tissue culture plate. 2. Incubate plates for at least 60 min at 37 °C in a tissue culture incubator (see Note 1). 3. Wash wells twice with sterile PBS. Aspirate the final wash before use.

3.2 Growth and TNFα-Mediated Activation of HUVECs

1. Starting with 100,000 HUVECs/well, culture the HUVECs on the gelatin-coated 48-well plates in a 2 mL/well of EGM-2 media for 48 h until the endothelial cells form a monolayer or 100% confluency. 2. Treat the endothelial cell monolayer with TNFα (vehicle vs. 2.5 ng/mL vs. 10 ng/mL) for 4 h in EBM-2 media containing 0.4% FBS (see Note 2).

3.3 Growth and Labeling of THP-1 Cells

1. Grow THP-1 cells in suspension culture in RPMI1640 media with 20% FBS until the desired number of cells for the assay (here one million cells) is reached. The culture medium is changed every other day until exponential growth phase is reached (see Note 3). 2. Pool all the THP-1 cell culture suspensions in one tube, count the cell concentration, and take a volume of THP-1 cell suspension containing one million cells. Spin down the cells at 300 g for 2 min, aspirate media, and resuspend the cells in 1 mL of the serum-free RPMI1640 media (106 cells/ml concentration). Add 2 μL of the 500× green fluorescent dye (see Note 4). 3. Incubate the THP-1 cells with the labeling dye for 60 min at 37 °C in a tissue culture incubator. Spin down cells at 300 g for 2 min, aspirate the medium, and wash cells with serum-free RPMI1640 media. Repeat the wash twice. Resuspend the cells in 10 mL RPMI1640 serum-free media (see Note 5).

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3.4 Assay of the Adhesion of the THP-1 Cells to the Endothelial Cell Monolayer

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1. Aspirate endothelial culture media and wash once with serumfree RPMI1640 media. Add the appropriate volume of the labeled THP-1 cells to each well already containing the HUVEC monolayer (see Note 6). In this case, add 400 μL of the THP-1 monocyte cell suspension to add 40,000 THP-1 cells (see Note 7). 2. Incubate the HUVEC-THP-1 mix for 1 h in a 37 °C tissue culture incubator. 3. Carefully discard or aspirate the media from each well without allowing wells to dry (see Note 8). Gently wash each well three times with 250 μL of diluted cell wash buffer to completely remove nonadherent THP-1 cells without dislodging the adherent cells. Leave the cell wash buffer of the last wash on the cells (see Note 8). 4. Very quickly take a picture of each well under an inverted fluorescence microscope to be used later for counting the adherent leukocytes (see Note 9). 5. Aspirate the final wash and add 150 μL of diluted cell lysis buffer to each well. Incubate 5 min at room temperature with shaking to ensure complete lysis of the adherent THP-1 cells. 6. Quickly transfer 100 μL from each well to wells of a 96-well plate suitable for fluorescence reading and read fluorescence with a fluorescence plate reader set at 480 nm excitation and 520 nm emission and read fluorescence intensity in a fluorescent plate reader. To validate the method, we determined the relationships between the fluorescence signal intensity and the number of labeled THP-1 cells and found a very nice linear relationship from THP-1 cells as low as 2500 cells to THP-1 cells as high as 50,000 (Fig. 2). To further validate the method, we quantified adhesion of the THP-1 cells to the HUVEC monolayer in response to activation of the endothelial cells with the pro-inflammatory factor TNFα. In agreement with compelling published data demonstrating that TNFα induces adhesion of inflammatory cells to the endothelium [12, 13], we found that the number of THP-1 adhering to the endothelial monolayer increased in response to TNFα treatment in a dose-dependent manner, based on the visualization of adherent cells under microscope (Fig. 3a), manual counting of the adherent cells (Fig. 3b), and the fluorescence signal intensity measure of the number of fluorescently labeled adherent THP-1 cells (Fig. 4).

4

Notes 1. Coat the plate with 0.1% gelatin for at least an hour. Extra plates can be stored in a refrigerator for a week.

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60000

y = 0.976x R² = 0.9901

50000

RFU

40000

30000

20000

10000

0

0

10000

20000

30000

40000

50000

60000

Cell Number

Fig. 2 Linearity of the fluorescence signal in response to increasing number of labeled THP-1 cells. Human monocytic THP-1 cells were labeled with the fluorescence dye as described in the Methods section, and an increasing number of labeled THP-1 cells were titrated in 75 μL of cell washing buffer and lysed with 75 μL of the lysis buffer. Fluorescence intensity was quantified as described in the Methods using a Tecan GENios Microplate Reader (Molecular Devices) with a 485/538 nm filter set and 530 nm cutoff. Fluorescence values are relative fluorescence units (RFU)

2. TNFα treatment is performed in EBM-2 media containing 0.4 FBS. To ensure consistency, determine the total volume of the media needed (2 mL/well) and add the desired amount of TNFα to the media and mix well before dispensing media in each well. In addition, before initiating the TNFα treatment, ensure that all the rich EGM-2 media is removed by washing cells at least once with PBS. 3. Because the THP-1 cells grow slower than the HUVECs, the growth of these cells is initiated earlier so that they are ready in sufficient number for the assay 2 days after the initiation of HUVEC growth. 4. Precise counting of the THP-1 cells to determine that the volume of TPH-1 culture suspension needed for labeling is critical to ensure consistency of labeling and of the adhesion assay.

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A Vehicle

TNFα (2.5 ng/ml)

TNFα (10 ng/ml)

B Number of aached THP1 monocytes

1400

*

1200 1000 800 *

600 400 200 0

vehicle

TNFa (2.5ng)

TNFa (10 ng)

Fig. 3 (a) TNFα dose-dependently increases human monocytic THP-1 cell adhesion to HUVEC monolayer as visualized by fluorescence microscopy. The HUVEC monolayer in a 48-well plate was treated with 0 or 2.5 or 10 ng/mL TNFα for 4 h. Fluorescently labeled THP-1 cells (40,000 cells/well) were allowed to attach to the HUVEC monolayer for 60 min, and nonadherent THP-1 cell was washed out, and the adherent cells were visualized and photographed under an inverted fluorescent microscope using a green filter (images were acquired using Olympus IX71 Inverted Microscope with Olympus micro DP 70 software, at 40× magnification). Presented are images taken from each of the three replicate wells and each is a representative image from three different fields taken per well. (b) TNFα dose-dependently increases human monocytic THP-1 cell adhesion to HUVEC monolayer as determined by manual counting of adherent THP-1 cells. HUVECs and THP-1 cells were processed as in Fig. 3a above, and the THP-1 cells adherent to the HUVEC monolayer were photographed under an inverted fluorescent microscope as above. Images of three different fields were taken per well and counted, and the numbers of adhering cells in each of the three fields were averaged as a semiquantitative measure of leukocyte recruitment per well/experimental unit, three wells per treatment. Bars are the standard errors of the mean

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*

Adhesion of THP-1 cells to HUVEC (RFU)

4750 4250 3750 3250 2750

*

2250 1750 1250 750

Vehicle

TNFa-2.5ng/ml

TNFa-10ng/ml

Fig. 4 TNFα dose-dependently increases human monocytic THP-1 cell adhesion to HUVEC monolayer as measured by fluorescence signal intensity. HUVECs and THP-1 cells were processed as described in Fig. 3a, and as a more sensitive quantitative measure of leukocyte adhesion, the THP-1 cells adherent to the HUVEC monolayer were lysed as described in the Methods section. The fluorescence intensity – a quantitative measure of the number of THP-1 cells adherent to the endothelium – was determined with the fluorescence plate reader as described in Fig. 2. At least three wells were used per experimental unit and bars represent standard errors of the mean

5. When preparing THP-1 monocytes, carefully aspirate the supernatant after centrifuging to avoid losing cells. 6. Adding appropriate number of THP-1 monocytes cells to each well of the HUVEC monolayer is critical. Adding too many cells will lead to nonspecific binding to the endothelial cells (see Fig. 2 and Note 3). 7. Ensure that the volume of labeled THP-1 cells covers all the endothelial monolayer. 8. When washing the cells, work fast to ensure that wells do not dry. 9. If unable to take a picture of the whole well to be able to determine the number of all adherent cells in the whole well, average at least three separate fields per well.

Acknowledgments This work was supported by NIH grants HL084725, HL003676, HL100258, and RR03034.

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References 1. Ley K, Laudanna C, Cybulsky MI, Nourshargh S (2007) Getting to the site of inflammation: the leukocyte adhesion cascade updated. Nat Rev Immunol 7(9):678–689 2. Zhong L, Simard MJ, Huot J (2018) Endothelial microRNA regulating the NF-κB pathway and cell adhesion molecules during inflammation. FASEB J 32:4070–4084 3. Al-Mehdi AB, Tozawa K, Fisher AB, Shientag L, Lee A, Muschel RJ (2000) Intravascular origin of metastasis from the proliferation of endothelium-attached tumor cells: a new model for metastasis. Nat Med 6:100–102 4. Leong HS, Robertson AE, Stoletov K, Leith SJ, Chin CA, Chien AE, Hague MN, Ablack A, Carmine-Simmen K, McPherson VA et al (2014) Invadopodia are required for cancer cell extravasation and are a therapeutic target for metastasis. Cell Rep 8:1558–1570 5. Kienast Y, von Baumgarten L, Fuhrmann M, Klinkert WE, Goldbrunner R, Herms J, Winkler F (2010) Real-time imaging reveals the single steps of brain metastasis formation. Nat Med 16:116–122 6. Cardoso TC, Pompeu TE, Silva CLM (2017) The P2Y(1) receptor-mediated leukocyte adhesion to endothelial cells is inhibited by melatonin. Purinergic Signal 13(3):331–338 7. Bendas G, Borsig L (2012) Cancer cell adhesion and metastasis: selectins, integrins, and the inhibitory potential of heparins. Int J Cell Biol 2012:676731, 1

8. Mitroulis I, Alexaki VI, Kourtzelis I, Ziogas A, Hajishengallis G, Chavakis T (2015) Leukocyte integrins: role in leukocyte recruitment and as therapeutic targets in inflammatory disease. Pharmacol Ther 147:123–135 9. Norman MU, Kubes P (2005) Therapeutic intervention in inflammatory diseases: a time and place for anti-adhesion therapy. Microcirculation 12(1):91–98 10. Weber GF (2013) Why does cancer therapy lack effective anti-metastasis drugs? Cancer Lett 328:207–211 11. Jandl K, Marsh LM, Hoffmann J, Mutgan AC, Baum O, Bloch W, ThekkekaraPuthenparampil H, Kolb D, Sinn K, Klepetko W, Heinemann A, Olschewski A, Olschewski H, Kwapiszewska G (2020) Basement membrane remodeling controls endothelial function in idiopathic pulmonary arterial hypertension. Am J Respir Cell Mol Biol 63(1):104–117 12. Zerr M, Hechler B, Freund M, Magnenat S, Lanois I, Cazenave JP, Le´on C, Gachet C (2011) Major contribution of the P2Y1 receptor in purinergic regulation of TNFα-induced vascular inflammation. Circulation 123(21): 2404–2413 13. Choi HJ, Kim NE, Kim BM, Seo M, Heo JH (2018) TNF-α-induced YAP/TAZ activity mediates leukocyte-endothelial adhesion by regulating VCAM1 expression in endothelial cells. Int J Mol Sci 19(11):3428

Chapter 19 Determination of Tight Junction Integrity in Brain Endothelial Cells Based on Tight Junction Protein Expression Himakarnika Alluri, Chander Sekhar Peddaboina, and Binu Tharakan Abstract Blood-brain barrier (BBB) dysfunction and hyperpermeability that occurs following traumatic and ischemic insults lead to various downstream ill effects such as cerebral edema and elevation of intracranial pressure. The inter-endothelial tight junctions that consist of tight junction proteins are critical regulators of BBB dysfunctions and hyperpermeability. The major tight junction-associated proteins of the BBB are occludin, claudins, and junctional adhesion molecules that are intracellularly linked to the adaptor protein zonula occludens-1 (ZO-1). Quantitative measurement of tight junction-associated proteins provides valuable insight into barrier integrity and mechanisms that regulate microvascular hyperpermeability. Western blot analysis is a commonly used method to separate and identify proteins in a mixture using gel electrophoresis. Understanding the changes in the expression of one or more of these proteins is critical to evaluating barrier integrity and permeability in health and disease. Furthermore, studying them will provide insight into the associated downstream signaling pathways and evaluation of therapeutic approaches for regulating BBB permeability. Herein, we have described the protocol for immunoblot analysis of ZO-1 as an indicator of tight junction integrity in brain microvascular endothelial cells. Key words Endothelial tight junctions, Tight junction proteins, Blood-brain barrier permeability, Zonula occludens-1

1

Introduction Blood-brain barrier (BBB) is a semipermeable membrane separating systemic circulation from the brain parenchyma. Microvascular hyperpermeability, an abnormal extravasation of plasma proteins and fluid into extravascular space in the brain, occurs at the blood-brain barrier. Alterations in BBB integrity following inflammation lead to vascular hyperpermeability and vasogenic edema [1, 2]. Optimal functioning of the BBB is crucial for maintaining the homeostasis of the brain. Inter-endothelial tight junctions of the BBB are the key structural and functional elements that play a major role in maintaining BBB integrity [3]. Tight junction

Binu Tharakan (ed.), Vascular Hyperpermeability: Methods and Protocols, Methods in Molecular Biology, vol. 2711, https://doi.org/10.1007/978-1-0716-3429-5_19, © Springer Science+Business Media, LLC, part of Springer Nature 2024

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proteins include transmembrane occludin, claudins, junctional adhesion molecules, and membrane-bound tight junction protein zonula occludes [ZO]. Zonula occludens-1 (ZO-1) are scaffolding molecules that link the transmembrane tight junctions and the actin cytoskeleton. ZO-1 plays an important role in maintaining the function of BBB including its permeability [3]. In vitro brain injury experimental studies have shown upregulation of IL-1β, a pro-inflammatory cytokine that can disrupt the BBB by deregulating the expression of tight junction proteins such as ZO-1 [4, 5]. ZO-1, a 220 kDa tight junction protein, can be measured to determine its role in determining tight junction integrity, by immunoblotting (Fig. 1). The current Western blot protocol outlines a detail technique on how the proteins are resolved over an SDS polyacrylamide gel and transferred to a nitrocellulose membrane to be detected by the ECL detection technique.

2

Materials 1. Rat brain microvascular endothelial cells (RBMVECs) [Cell Applications Inc., San Diego, CA]. 2. RBMVEC Media [Cell Applications Inc., San Diego, CA]. 3. Fibronectin [Sigma Aldrich, St. Louis, MO]. 4. Interleukin-1β, human [Sigma Aldrich, St. Louis, MO]. 5. ZO-1 Monoclonal Antibody [Thermo Fisher Scientific, Carlsbad, CA]. 6. Opti-MEM (1×)/reduced serum medium [Thermo Fisher Scientific, Carlsbad, CA]. 7. HyClone Dulbecco’s phosphate-buffered saline (PBS, without calcium, magnesium, or Phenol Red) [Thermo Fisher Scientific, Carlsbad, CA]. 8. Anti-rabbit IgG-FITC secondary antibody [Santa Cruz Biotechnology, Inc. Santa Cruz, CA]. 9. NuPAGE Novex® 10% Bis-Tris protein gels [Thermo Fisher Scientific, Carlsbad, CA]. 10. NuPAGE® MOPS SDS Running Buffer [Thermo Fisher Scientific, Carlsbad, CA]. 11. RIPA cell lysis buffer [Thermo Fisher Scientific, Carlsbad, CA]. 12. NuPAGE® Transfer Buffer [Thermo Fisher Scientific, Carlsbad, CA]. 13. Nitrocellulose/filter paper sandwich [Thermo Fisher Scientific, Carlsbad, CA]. 14. Protease inhibitor cocktail (100×) [Thermo Fisher Scientific, Carlsbad, CA].

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Fig. 1 Western blot analysis of zonula occludens-1 (ZO-1) from rat brain microvascular endothelial cell (RBMEC) lysates. RBMECs were exposed to increasing concentrations of hydrogen peroxide (H2O2) followed by immunoblot analysis. H2O2 treatment showed no significant effect on ZO-1 protein expression at the concentrations tested (ZO-1 was normalized to β-actin expression). This data was taken from a previously published article from our lab (Anasooya Shaji et al., Scientific Reports, 2019, 9.133). Creative Commons License https://creativecommons.org/licenses/)

15. Pierce™ ECL Plus Western Blotting Substrate [Thermo Fisher Scientific, Carlsbad, CA]. 16. Goat anti-mouse IgG-HRP [Santa Cruz Biotechnology, Inc. Santa Cruz, CA]. 17. CL-XPosure™ Film [Thermo Fisher Scientific, Carlsbad, CA].

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Methods

3.1 RBMEC Cell Lysate Preparation

1. Rat brain microvascular endothelial cells (RBMVECs) are cultured in a 10 cm Petri dish coated with 5% fibronectin (50 μg/ml in PBS). 2. Once cells reach 80% confluency, remove the culture media and incubate the cells in reduced serum medium. 3. Wash cells twice with ice-cold PBS and incubate in 1 mL of ice-cold RIPA cell lysis buffer (1×) along with protease inhibitor cocktail (1×) for 5 min. 4. Use a cell scraper to dissociate the cells from the Petri dish. Collect the soup into a 1.5 mL microcentrifuge tube and sonicate for 30 s on ice to fragment the cells. 5. Centrifuge the lysate at 14,000g for 10 min at 4 °C. 6. Carefully collect the supernatant in a fresh centrifuge tube, and determine protein concentration using BCA protein assay kit.

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3.2 Western Blot Analysis of ZO-1

1. Prepare equal amounts of total protein (50 G) in loading buffer and add β-mercaptoethanol to a final concentration of 5% (v/v %), heat at 95 °C for 5 min on a heating block and transfer onto the ice, and prepare SDS-PAGE setup. 2. Load protein ladder (5 μL). 3. Proteins are separated by sodium dodecyl sulfatepolyacrylamide gel electrophoresis (SDS-PAGE) on 10% Bis-Tris precast gels at constant voltage (120 V) for 90 min in MOPS SDS running buffer. 4. Open the gel plates using a spatula, and immerse the gel in DI water to remove any traces of running buffer. 5. Add 200 mL of methanol to 800 mL of 1× NuPAGE® Transfer Buffer to prepare protein transfer buffer. 6. Soak nitrocellulose membrane and filter paper in transfer buffer to equilibrate, transfer the SDS-gel on to the filter paper, and gently place the nitrocellulose membrane on top followed by a second filter paper on top of it. 7. Create a sandwich by placing the filter-nitrocellulose-gel-filter paper between two blotting sponge pads and transfer it into blotting cassette. Insert the blotting cassette into protein transfer setup with the proper orientation. 8. Proteins are transferred onto the nitrocellulose membrane at constant voltage (30 V) overnight. 9. On the following day, remove the membrane from the transfer setup, and block with 5% nonfat dry milk in Tris-buffered saline (TBS) with 0.05% Tween-20 (blocking buffer) for 1 h. 10. Incubate the membrane with primary mouse monoclonal anti ZO-1 antibody diluted at 1:250 in 5% nonfat dry milk TBS-T overnight at 4 °C. 11. On the next day, wash the membrane three times with TBS-T. 12. Block the membrane in 5% nonfat dry milk in Tris-buffered saline (TBS) with 0.05% Tween-20 for 1 h at room temperature. 13. Incubate the membrane with the goat anti-mouse IgG-HRP conjugated secondary antibody diluted at 1:2000 in 5% nonfat dry milk in Tris-buffered saline (TBS) with 0.05% Tween-20 for 2 h at room temperature. 14. Wash the membrane in TBS-T with wash buffer replaced every 15 min for a total of three washes. 15. Prepare fresh ECL Western blotting substrate and incubate the membrane in the substrate for 1 min at room temperature.

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16. Pick the membrane and dab the edges to remove excess solution, cover it in a plastic wrap, and use a roller to remove any trapped bubbles before placing in the film cassette. 17. Expose the membrane to an X-ray film and develop using an autoradiogram. 18. Developed X-ray films are scanned and the band intensity is quantified using ImageJ software.

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Notes 1. During the protein extraction process from the cells, the sonication step produces excess heat which can affect the proteins and so the process must be done on ice. 2. Fill empty wells with loading buffer so that all the wells run equally and make sure to use the right molecular weight protein ladder that is compatible with the gel and the running buffer. 3. While running the acrylamide gel, running buffer gets hot overtime at high voltages, do not run the gel for too long at high voltage (140 V+) as this will damage the gel. 4. Since the transfer buffer contains 20% methanol, do not use the old buffer; always prepare fresh transfer buffer. Old transfer buffer can be used for initial steps where the nitrocellulose membrane and the filter papers need to be soaked before setting up the transfer cassette. 5. Protein transfer efficiency onto nitrocellulose membrane from the acrylamide gel is better when the transfer buffer is maintained at or close to 4 °C, use either ice packs or run the transfer in a 4 °C refrigerator overnight at low voltage. 6. The membrane should not dry during any of the primary and secondary antibody incubation, washing steps. 7. All membrane washes in TBS-T and the secondary antibody incubation should be done at room temperature. 8. ECL Western blot substrate should be used within 1 h of preparation. The substrate should be prepared right before the final membrane wash postsecondary antibody incubation. 9. Before the membrane is placed in the cassette, all the trapped bubbles on the membrane should be removed using a roller to avoid poor signals during exposure. 10. The membrane should be exposed to the X-ray film in the dark, and the exposure time can be adjusted based on the signal intensity.

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11. Sample loading can be verified by assessing β-actin protein expression. 12. Use untreated cells to serve as controls.

Acknowledgments The authors acknowledge the support from the National Institutes of Health (NIH) grant 5 SC3 NS127765-02 (BT). References 1. Bolton SJ, Anthony DC, Perry VH (1998) Loss of the tight junction proteins occludin and zonula occludens-1 from cerebral vascular endothelium during neutrophil-induced blood-brain barrier breakdown in vivo. Neuroscience 86(4): 1245–1257 2. Rigor RR et al (2012) Interleukin-1betainduced barrier dysfunction is signaled through PKC-theta in human brain microvascular endothelium. Am J Physiol Cell Physiol 302(10): C1513–C1522

3. Alluri H et al (2015) Blood-brain barrier dysfunction following traumatic brain injury. Metab Brain Dis 30(5):1093–1104 4. Chen F et al (2009) Disruptions of occludin and claudin-5 in brain endothelial cells in vitro and in brains of mice with acute liver failure. Hepatology 50(6):1914–1923 5. Wu B et al (2010) Ac-YVAD-CMK decreases blood-brain barrier degradation by inhibiting caspase-1 activation of interleukin-1beta in intracerebral hemorrhage mouse model. Transl Stroke Res 1(1):57–64

Chapter 20 Evaluation of Glycolysis and Mitochondrial Function in Endothelial Cells Using the Seahorse Analyzer Zeinab Y. Motawe, Salma S. Abdelmaboud, and Jerome W. Breslin Abstract Endothelial bioenergetics have emerged as a key regulator of endothelial barrier function. Glycolytic parameters have been linked to barrier enhancement, and interruption with mitochondrial complexes was shown to disrupt endothelial barrier. Therefore, a new technology that has been introduced to assess bioenergetics and metabolism has also made it possible to determine roles of specific energy production pathways in endothelial health. The Seahorse extracellular flux analysis by Agilent technologies is a state of the art tool that has been more frequently used to evaluate bioenergetics of endothelial cells. This chapter includes details about different assays that can be used to study endothelial cells using the Seahorse analyzer and how interpretation of the results can provide novel insight about endothelial metabolism. Key words Glycolysis, ATP rate, Endothelium, Seahorse, Metabolism, Endothelial barrier, Mitochondria

1

Introduction Metabolism is broadly defined as the sum of biochemical processes in living organisms that either produce or consume energy [1] and can be divided into two processes: anabolism and catabolism. Anabolic pathways require energy to make macromolecules such as lipid and nucleotides, whereas the catabolic pathway breaks molecules to generate energy [2]. The main energy production pathways are mitochondrial oxidative phosphorylation and anaerobic glycolysis. Mitochondrial energy production is the main energy production pathway in many cell types such as cardiac myocytes of the heart or hepatocytes of the liver [3, 4]. In endothelium, anaerobic glycolysis has been shown to be the main energy production pathway [5]. Oxygen consumption is required for energy production in cells [6–8], and the oxygen consumption rate has been determined to be a reliable indicator of active cellular metabolism and mitochondrial function [9–11].

Binu Tharakan (ed.), Vascular Hyperpermeability: Methods and Protocols, Methods in Molecular Biology, vol. 2711, https://doi.org/10.1007/978-1-0716-3429-5_20, © Springer Science+Business Media, LLC, part of Springer Nature 2024

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Several methods have been developed to measure OCR. These include an oxygen electrode, a specialized piece of equipment that typically yields low sample throughput [12], a phosphorescent oxygen probe that has been effective for measuring oxygen consumption rates in whole cells [13], and fluorescent detection of OCR which was also previously used [14]. Agilent technologies introduced a new label-free, high-throughput method of measuring both OCR and the extracellular acidification rate (ECAR), another cellular energy production indicator [15], in the same assay which is label-free, real time, and automatic in 2006. Since then, this technology has been used in over 5000 publications due to its reliability and reproducibility. Assay protocols can be tailored to target different tissues and cell types. Several standardized assays are commercially available, such as the Cell Mito stress test, glycolytic rate assay, glycolytic stress rest, and ATP rate assay kits. Endothelial cells rely on anaerobic glycolysis as their primary energy source, with 99% of glucose converted anaerobically to lactate and only 0.04% undergoing oxidative phosphorylation [5, 16]. This enables ATP generation in peripheral regions of the cell, in close proximity to cell junctions as well as lamellipodia and filopodia, and sparing the need to transfer ATP from more centrallocated mitochondria [17, 18]. In addition, glycolysis regulatory enzymes have been shown to colocalize with areas near cell junctions [19]. Active spatiotemporal patterns of ATP-dependent signaling were also shown at the cell periphery when cells were stimulated to enhance their barrier function [20]. Therefore, emerging methods to assess endothelial glycolysis with respect to pathobiology are needed to allow for novel discoveries to protect endothelial health pertaining to endothelial energy production. Despite the fact that mitochondria are not the main energy source in endothelial cells, and only occupy 2–6% of endothelial cellular volume, in contrast to 28% in hepatocytes and 32% in cardiac myocytes, they have important signaling roles in endothelium [21, 22]. Mitochondria are an important source of reactive oxygen species [23]. In addition, mitochondria play an important role in calcium homeostasis in endothelium which is critical for some endothelial functions such as NO production [24]. Moreover, endothelial mitochondria participate in the regulation of apoptosis [25]. There are many disease conditions that are characterized by endothelial mitochondrial dysfunction, such as atherosclerosis, diabetes, or hypertension [26–28]. Mitochondrial dysfunction was also shown in hemorrhagic shock and resuscitation in rats [29]. Mitochondrial complex III inhibition was shown to disrupt rat intestinal endothelial barrier function [29]. In addition, release of mitochondrial cytochrome c in the cytoplasm and activation of downstream effector caspase-3 was shown to lead to loss of

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endothelial cell barrier integrity in a hemorrhagic shock model [30]. Consequently, novel ways to assess mitochondrial function in endothelium are needed to expand the current knowledge and introduce new therapeutics. Seahorse technology allows for rapid assessment of mitochondrial function and glycolytic metabolism in endothelial cells with easy-to-use, reproducible assays. Different assays such as cell the Mito stress test, glycolytic stress test, glycolytic rate assay, or ATP rate assay have been used to assess endothelial cell functions in angiogenesis, NO production, coronary dysfunction, shear stress, and barrier function [31–36], with recently published examples shown in Table 1. This chapter details three of the Seahorse assays, namely, the Mito stress test, glycolytic rate assay, and ATP production rate assay. Also included is an introduction to Wave software which can be used to view and perform analysis of data obtained from the Seahorse analyzer. Wave is free to download from the Agilent Technologies website. 1.1 Cell Mito Stress Test

The Agilent Seahorse XFp Cell Mito stress test measures important parameters of mitochondrial function using real-time measurement of cellular OCR before and after addition of compounds that modulate certain steps in the mitochondrial transport chain [37], as shown in Fig. 1. The three compounds used in the assay are oligomycin, carbonyl cyanide-4 (trifluoromethoxy) phenylhydrazone (FCCP), and rotenone and antimycin. Oligomycin is injected first which inhibits ATP synthase (complex V). It decreases electron flow through the electron transport chain, reducing OCR [47]. FCCP is injected second and is an uncoupling agent that collapses the proton gradient and disrupts the mitochondrial membrane potential, resulting in uninhibition of the electron transport chain, allowing for oxygen consumption by complex IV to reach its maximum [48]. Lastly, the third injection is a mixture of the complex I inhibitor rotenone [49] and the complex III inhibitor antimycin A [50]. In combination, these compounds shut down mitochondrial respiration. The OCR measured under each of these conditions provides information of each of the parameters shown in Fig. 1.

1.2 Glycolytic Rate Assay

The Seahorse XFp glycolytic rate assay provides precise measurements of glycolytic rates for basal conditions and compensatory glycolysis following mitochondrial inhibition. This assay uses both ECAR and OCR measurements to define the glycolytic proton efflux rate (glycoPER) of the cells [51]. It is well established that glucose in cells is converted to pyruvate and then converted to either lactate in the cytoplasm or to CO2 and water in the mitochondria [52]. The conversion of glucose to lactate promotes the release of protons into the extracellular medium to maintain intracellular pH homeostasis [53]. This extracellular acidification is

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Table 1 Examples of endothelial cell assessment with Seahorse analysis Type of cells

Seahorse model

Number of cells/ well Assay used

XF96

40,000

Purpose

Authors

Assess tip cell metabolic behavior

Yetkin-Arik et al. [31]

1

Human umbilical vein endothelial cells (HUVEC) and dermal human microvascular endothelial cells (HMVEC)

2

Bovine aortic endothelial cells XF24 (BAEC)

20,000–60,000 Mito stress test

Assess mitochondrial reserve capacity in response to NO and ROS

Dranka et al. [32]

3

Human cardiac microvascular XF24 endothelial cells (HCMVEC)

Not specified

Mito stress test

Assess mitochondrial function in hyperglycemia

Joshi et al. [34]

4

BAEC

XF24

5000–10,000

Mito stress test

Assess effect of glucose on endothelial bioenergetics

Fink et al. [33]

5

HUVEC

XF24

10,000–80,000 Mito stress test

Study effects of atorvastatin on mitochondrial energy metabolism

Chang et al. [37]

6

HUVEC

XF24

20,000

Mito stress test

Study the effects of Li et al. [38] advanced glycation end products on mitochondrial energy

7

HUVEC

XF96

60,000

Glycolytic stress Study effects of test laminar shear stress on endothelial metabolism

Doddaballapur et al. [35]

8

Human aortic endothelial cells (HAEC)

XF96

30,000

Mito stress test

Study mitochondrial ROS and endothelial activation

Li [39]

9

HUVEC

XF24

30,000

Mito stress test

Study effects of ALDH2 on mitochondrial oxygen reserve capacity in ECs

Nanelli et al. [40]

10 HUVEC

XF24

62,000

Mito stress test

Study effects of VEGF on endothelial metabolism

Domigan et al. [41]

Mito fuel flex test, glycolytic stress test

(continued)

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Table 1 (continued) Seahorse model

Number of cells/ well Assay used

11 Cerebral vascular endothelial cells

XF96e

16,000

Mito stress test

12 Human corneal endothelial cells

XFe24

Not specified

Mito stress test, Mitochondrial glycolytic changes in DM stress test

13 Pulmonary arterial endothelial cells

XF24

75,000

Mito stress test, Effects of Sun et al. [44] glycolytic endothelin 1 on stress test endothelial bioenergetics

14 HUVEC

XFp

40,000

Glycolytic rate assay, ATP production rate assay

15 HUVEC

XF24

60,000

Glycolytic stress Effects of NRF2 test on endothelial glycolysis

Kousmanen et al. [45]

16 Isolated mouse brain microvessel suspension

XF24

Not specified

Mito stress test

Sure et al. [46]

Type of cells

Purpose

Authors

Mitochondrial Hu et al. [42] function in cerebrovascular endothelial cells in stroke

Effects of σ1 activation on endothelial bioenergetics

Aldrich et al. [43]

Motawe et al. [36]

measured in real time and give an indication about glycolytic proton efflux rate. During the assay, Rot/AA (inhibitors of mitochondrial electron transport chain) are injected to inhibit mitochondrial oxygen consumption (and consequently CO2-derived proton efflux). The second injection is 2-deoxy-D-glucose (2-DG), which inhibits glycolysis through competitive binding of glucose hexokinase [54]. Measurement of OCR and ECAR under these conditions provides information about the parameters shown in Fig. 2. 1.3 ATP Production Rate Assay

The Seahorse ATP production rate assay allows for differentiation whether ATP is predominantly produced by mitochondrial oxidative phosphorylation (OXPHOS) or glycolysis. This assay uses oligomycin and the mixture of rotenone and antimycin A that allows the calculation of the mitochondrial and glycolytic realtime ATP production rates in live cells [55]. An example of the type of data that can be obtained with this protocol is shown in Fig. 3.

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Fig. 1 Seahorse XFp Cell Mito stress test assay profile showing the parameters of mitochondrial function. The ATP synthase inhibitor oligomycin is injected first and decreases electron flow through the electron transport chain, reducing OCR. FCCP is injected second which collapses the proton gradient and disrupts the mitochondrial membrane potential, resulting in uninhibition of the electron transport chain, allowing for oxygen consumption by complex IV to reach its maximum. Lastly, the third injection is a mixture of the complex I inhibitor rotenone and the complex III inhibitor antimycin A. In combination, these compounds shut down mitochondrial respiration. Nonmitochondrial oxygen consumption = minimum rate measurement after rotenone/antimycin A injection. Basal respiration = (last rate measurement before first injection) - (non-mitochondrial respiration rate). Maximal respiration = (maximum rate measurement after FCCP injection) (non-mitochondrial respiration) H+ (proton) leak = (minimum rate measurement after oligomycin injection) - (non-mitochondrial respiration). ATP production = (last rate measurement before oligomycin injection) - (minimum rate measurement after oligomycin injection). Spare respiratory capacity = (maximal respiration) - (basal respiration). (Reproduced with permission copyright from Agilent technologies)

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Materials 1. Seahorse analyzer. 2. Non-CO2 Incubator. 3. Seahorse cell culture miniplates. 4. Seahorse sensor cartridges.

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Fig. 2 Agilent Seahorse XFp glycolytic rate assay profile. Inhibition of mitochondrial function by rotenone and antimycin A (Rot/AA) enables calculation of mitochondrial-associated acidification. Subtraction of mitochondrial acidification to total proton efflux rate results in glycolytic proton efflux rate (glycoPER). Glycolytic proton efflux rate (glycoPER) represents the proton efflux rate produced from glycolysis (discounting the effect of CO2dependent acidification). Compensatory glycolysis represents the rate of glycolysis in cells after the addition of Rot/AA, inhibiting oxidative phosphorylation and leading the cells to compensatory upregulate glycolysis to meet the cells’ energy demands. Post-2-DG acidification represents other sources of extracellular acidification that are not linked to glycolysis or mitochondrial TCA activity as well as any residual glycolysis not fully inhibited by 2-DG. (Reproduced with permission copyright from Agilent technologies)

5. Sterile water. 6. Calibrant solution (Agilent). 7. Seahorse DMEM media (assay medium, Agilent). 8. Glucose solution (1.0 mM, Agilent). 9. L-Glutamine solution (200 mM, Agilent). 10. Sodium pyruvate solution (100 mM, Agilent). 11. Cell culture supplies: trypsin, culture media, PBS. 12. Materials for Cell Mito stress test: oligomycin, FCCP, rotenone/antimycin A (Rot/AA) mixture (included in the Seahorse Cell Mito stress test kit).

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Fig. 3 Representation of Agilent Seahorse XFp Real-Time ATP rate assay kinetic profile of OCR and ECAR measurements. Basal OCR and ECAR rates are first measured. Injection of oligomycin results in an inhibition of mitochondrial ATP synthesis that causes a decrease in OCR, allowing the mito-ATP production rate to be quantified. ECAR data allows calculation of total proton efflux rate (PER). Inhibition of mitochondrial respiration with rotenone plus antimycin A allows accounting for mitochondrial-associated acidification, and when combined with PER data allows calculation of the glycoATP production rate. (Reproduced with permission copyright from Agilent technologies)

13. Materials for glycolytic rate assay: 2-deoxy glucose (2-DG), Rot/AA (included in Seahorse glycolytic rate assay kit). 14. Materials for ATP rate assay: oligomycin, Rot/AA (included in Seahorse ATP rate assay kit). 15. 10× RIPA buffer. 16. BCA protein assay kit. 17. Plate reader for the BCA protein assay.

3

Methods We will describe three commonly used Seahorse assays that have been used to study endothelial bioenergetics, namely, the Cell Mito stress test, glycolytic rate assay, and ATP rate assay. We will use the Seahorse XFp as an example, which features eight assay wells per experiment. All of the methods generally follow the Agilent Seahorse instruction manuals, although some minor modifications were made to optimize results. Some modifications of the protocols may also be needed if using the Seahorse XF24 or XF96 models, which feature 24 and 96 assay wells, respectively.

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The initial preparation and seeding of cells into the assay cultureware is typically done 1 day before the assay. However, in some instances, such as when the development of mature junctions between endothelial cells is desired, or if the cells have been transfected and time is needed for optimal gene expression or knockdown, the seeding may be done several days prior to the assay. Seahorse XFp miniplates contain eight wells marked with the letters A through H. It is important to note that in general, wells B through G are only seeded with cells to be assayed, whereas wells A and H are left without cells and used as background wells. 3.1 Preparation of Cells and Cartridges

1. On the day prior to the Seahorse assay, the Seahorse analyzer should be turned on and left on overnight. Add 400 μL of sterile water or PBS to the moats around the cell culture wells. 2. Coat all eight wells with 1.5% porcine gelatin by adding 50 μL of gelatin to each well, shaking the plate to make sure that gelatin is covering the entire surface on which the cells will grow, and then aspirate any excess gelatin solution from the well. 3. Add 200 μL of growth medium only (no cells) to wells A and H. 4. Choose the desired seeding concentration. Cell seeding density can vary according to the purpose of the study, the type of endothelial cells used, and the surface area of the well, which depends on the type of Seahorse analyzer used. Refer to Table 1 for examples of seeding densities recently used in literature for different endothelial cell types. 5. Harvest the cells using standard procedures. Resuspend the cells in appropriate growth medium, count, and then dilute to the desired seeding concentration. 6. Add 200 μL of the cell suspension to wells B through G. 7. Allow the cells to grow overnight (or longer if desired) in a cell culture incubator. For cells being cultured for longer periods, ensure that the moats do not dry out. 8. Check the growth and health of cells using a microscope daily.

3.2 Hydration of Sensor Cartridges

1. One day prior to performing the assay, add 200 μL of sterile water to each of the eight wells of a sensor cartridge and 400 μL to each moat. Leave overnight in a non-CO2 incubator at 37 ° C. 2. On the day of the assay, change the sterile water to calibrant solution at least 2 h before the start of the assay. The hydrated cartridges need to be placed in a non-CO2 incubator at 37 °C until the assay is performed.

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3.3 Assay Media Preparation

1. On the day of the assay, warm the L-glutamine solution in a warm bead or water bath until it looks clear. 2. In a biosafety cabinet and under sterile conditions, add appropriate amounts of the glucose, glutamine, and pyruvate solutions to the desired amount of assay media (Seahorse DMEM) to a final concentration of glucose 10 mM, pyruvate 1 mM, and glutamine 2 mM. 3. Warm media in a 37 °C, non-CO2 incubator before the assay.

3.4 Mito Stress Test Instructions

1. Remove one foil pouch and the decapper tool from a Seahorse XFp Cell Mito stress test kit box.

3.4.1 Prepare Compound Stock Solutions and Working Solutions

2. Open the pouch and remove the three tubes containing oligomycin (blue cap), FCCP (yellow cap), and rotenone/antimycin A (red cap). Each vial should be tapped down to make sure that the contents are in the bottom of each vial before opening. 3. Remove the cap of each tube using the provided decapper tool. 4. Resuspend the contents of each tube with prepared assay medium in volumes by pipetting up and down (~10 times) to solubilize the compounds. The volumes of media to be added are provided in the kit instructions. After resuspension, these are the compound stock solutions. 5. Use the compound stock solutions to make compound working solutions for loading into the injection ports on sensor cartridges. For oligomycin, 1.5 μM is recommended for most cell types, while for Rot/AA, 0.5 μM is recommended, and for FCCP 1 μM is recommended. 6. Load the solutions into the appropriate ports on sensor cartridge. If performing the standard assay, only the modulators included with the kit (oligomycin, FCCP, and Rot/AA) will be added, and these can be loaded into ports A, B, and C, respectively. If modifying the assay to observe if a test compound acutely alters mitochondrial function, then that compound should be loaded into port A and then the modulators from the kit into ports B, C, and D. Each port is designed to deliver at a 1:10 dilution, so if each well is loaded with 180 μL of assay medium (see next subsection), then port A should be loaded at a volume of 20 μL, port B at 22 μL, port C at 25 μL, and port C at 27 μL to account for changes in volume with addition of each solution during the course of the assay.

3.4.2 Prepare Seahorse XFp Cell Culture Miniplate for Assay

1. Remove cell culture miniplates from a 37 °C CO2 incubator, and inspect the cells under a microscope to confirm confluence. 2. Remove the assay medium from water bath. 3. In a biosafety cabinet, change the cell culture growth medium in the cell culture miniplate to warmed assay medium and place

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the cell culture miniplate into a 37 °C non-CO2 incubator for 45 min to 1 h prior to the assay. Starting assay medium volume for cell plate is 180 μL per well. 3.4.3

Running the Assay

1. To run the Agilent Seahorse XFp Cell Mito stress test, click Start on the Seahorse analyzer, and select the Agilent Seahorse Cell Mito stress test default template. Click the right arrow on the Groups page, and then on the Protocol page click Start Assay. 2. Place the utility plate with the loaded assay cartridge on the instrument tray and click Continue. The instrument will perform a calibration that takes roughly 20 min. 3. When calibration has been completed, remove the utility plate and place the Seahorse XFp Cell Culture Miniplate on the tray, and press continue to start the assay.

3.5 Glycolytic Rate Assay

1. The same general procedure used for the Cell Mito stress test can be applied with glycolytic rate assay with some key differences. 2. Prepare stock compounds 2-DG (500 mM), Rot/AA (25 μM). 3. Dilute compounds to final concentration of 2-DG (50 Mm), Rot/AA (0.5 μM). 4. Load sensor cartridges (20 μL of Rot/AA to port A, 22 μL of 2-DG to port B). 5. Run assay, choosing the glycolytic rate assay option.

3.6

ATP Rate Assay

1. The procedures for this assay are largely the same as for the Cell Mito stress test, but some key differences: 2. Prepare stock compound solutions of oligomycin (75 μM) and Rot/AA (25 μM). 3. Dilute compounds to a final concentration of oligomycin (1.5 μM), Rot/AA (0.5 μM). 4. Load sensor cartridges (20 μL of oligomycin to port A, 22 μL of Rot/AA to port B). 5. Run assay, but choose the ATP rate assay option.

3.7 Protein Assay for Normalization

1. After the assay is completed, the protein content of each well can be determined as a way to minimize variation due to any differences in cellular mass between wells. Lysis of the cells with RIPA lysis buffer and BCA protein assay or other standard lysis/extraction buffers and protein assays are suitable for this purpose.

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2. Before the Seahorse assay has concluded, prepare RIPA buffer from 10× RIPA, 100× HALT protease and phosphatase inhibitor, and water, and chill on ice. 3. After the Seahorse assay has concluded and the assay plate is removed, place the assay plate on ice (or an ice brick). 4. Remove all media from all wells and wash twice with ice-cold PBS. 5. Remove PBS and add 50 μL of cold RIPA buffer to each well for an 8-well assay plate. Wells A and H, which contain no cells, will serve as background controls for the Seahorse assay. 6. Leave the assay plate containing RIPA buffer on ice for 20 min for protein extraction. 7. Prepare the BCA assay reagent by mixing components A and B according to the instructions on the bottles in a ratio of 50:1 respectively. 8. Set up a 96-well plate with protein standards to generate a standard curve, plus duplicate wells for the unknowns that will come from the Seahorse assay plate. 9. Add 10 μL of each standard and sample to the 96-well BCA assay plate. 10. Add 200 μL of the BCA working reagent to each well. 11. Incubate the BCA assay plate in a 37 °C incubator for 30 min. 12. Take the plate to a plate reader to read absorbance values at 562 nm. 13. Generate a standard curve with the BCA standards and use the equation to calculate the concentration of the samples in μg/μL. 14. Multiply the values by 50 to get the amount of protein in each sample well. These values will be used to normalize using Wave software. 3.8

Data Analysis

1. At the conclusion of each experiment, data can be extracted from the Seahorse analyzer in Excel, GraphPad Prism, or Wave files. 2. Using Wave software, data can be extracted as report generators. 3. The Agilent Seahorse XF Cell Mito stress test, glycolytic rate, or ATP production rate assay report generators automatically calculate the Agilent Seahorse XFp assay parameters from Wave data that has been exported to Excel. 4. In Wave home page, press the tab “normalize” which will lead you to a page in which you can insert your normalization data. Specify whether you are using number of cells or amount of

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proteins in normalization. Enter protein quantity in each well and click Apply. This will normalize the seahorse data. You can check or uncheck the tab “normalize” in the Wave home page to compare normalized data to nonnormalized data. Combining Data from Multiple Assays 1. After doing the multiple experiments to verify reproducibility, the data can be combined for analysis in software such as GraphPad Prism. 2. After running each experiment, the data for that experiment can be saved from the Seahorse instrument as a Prism or Excel file. 3. Alternatively, Wave software can also be used to convert the data extracted into a Prism file. Data from separate experiments can be copied and pasted between prism files in order to make comparisons with the larger dataset. 4. Assay readouts describing mitochondrial or glycolytic parameters can also be combined using the Excel report generator that can be extracted through Wave software. It provides assay parameters for each well that can be copied to Prism files for statistical comparisons.

4

Troubleshooting 1. To overcome cell detachment during media changes, it is recommended that the media be changed slowly and with extra care not to touch the cells while pipetting. 2. Careful pipetting is critical for optimal results and minimizing variation between wells receiving the same treatments. 3. In case certain compounds don’t seem to provide the optimum response, double-check the storage conditions of the compounds and the optimum concentration for each compound. 4. Number of cells greatly affects the results. Normalization of the results in the number of cells or the total protein by doing a protein assay is strongly recommended to overcome variations in the data. 5. Wave software provided by Agilent technologies should be used for analysis. 6. Not seeding enough cells can lower the signal received in the assay, and also potentially facilitate cell detachment during media changes. 7. In case negative values are obtained, it is important to make sure not to have any cells in the background wells. In addition,

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double-check the expiration date of the cartridges (should be done prior to starting the assay). Hydrating the cartridges in water overnight and in calibrant solution prior to the assay is absolutely necessary for the assay to work properly. 8. It is recommended to keep the seahorse analyzer always on to avoid accidently forgetting to turn it on the day before the assay. 9. In a rare situation where the instrument is not showing that it is at 37 °C, it is recommended to restart the instrument. This should bring the temperature back to 37 °C if the instrument is already set to this set point. References 1. DeBerardinis RJ, Thompson CB (2012) Cellular metabolism and disease: what do metabolic outliers teach us? Cell 148(6):1132–1144 2. Schwartsburd PM (2017) Catabolic and anabolic faces of insulin resistance and their disorders: a new insight into circadian control of metabolic disorders leading to diabetes. Future Sci OA 3(3):FSO201 3. Bround MJ, Wambolt R, Luciani DS, Kulpa JE, Rodrigues B, Brownsey RW et al (2013) Cardiomyocyte ATP production, metabolic flexibility, and survival require calcium flux through cardiac ryanodine receptors in vivo. J Biol Chem 288(26):18975–18986 4. Rui L (2014) Energy metabolism in the liver. Compr Physiol 4(1):177–197 5. Zecchin A, Kalucka J, Dubois C, Carmeliet P (2017) How endothelial cells adapt their metabolism to form vessels in tumors. Front Immunol 8:1750 6. Nakazawa MS, Keith B, Simon MC (2016) Oxygen availability and metabolic adaptations. Nat Rev Cancer 16(10):663–673 7. Wilson DF, Erecinska M (1986) The oxygen dependence of cellular energy metabolism. Adv Exp Med Biol 194:229–239 8. Wilson DF, Erecinska M (1985) Effect of oxygen concentration on cellular metabolism. Chest 88(4 Suppl):229S–232S 9. Salin K, Auer SK, Rey B, Selman C, Metcalfe NB (2015) Variation in the link between oxygen consumption and ATP production, and its relevance for animal performance. Proc Biol Sci 282(1812):20151028 10. Elliott JM, Davison W (1975) Energy equivalents of oxygen consumption in animal energetics. Oecologia 19(3):195–201 11. Decleer M, Jovanovic J, Vakula A, Udovicki B, Agoua REK, Madder A et al (2018) Oxygen

consumption rate analysis of mitochondrial dysfunction caused by Bacillus cereus cereulide in Caco-2 and HepG2 cells. Toxins (Basel) 10(7):266 12. Clark LC Jr, Wolf R, Granger D, Taylor Z (1953) Continuous recording of blood oxygen tensions by polarography. J Appl Physiol 6(3): 189–193 13. Will Y, Hynes J, Ogurtsov VI, Papkovsky DB (2006) Analysis of mitochondrial function using phosphorescent oxygen-sensitive probes. Nat Protoc 1(6):2563–2572 14. Yun CW, Han YS, Lee SH (2019) PGC-1alpha controls mitochondrial biogenesis in drugresistant colorectal cancer cells by regulating endoplasmic reticulum stress. Int J Mol Sci 20(7):1707 15. Zhang J, Nuebel E, Wisidagama DR, Setoguchi K, Hong JS, Van Horn CM et al (2012) Measuring energy metabolism in cultured cells, including human pluripotent stem cells and differentiated cells. Nat Protoc 7(6):1068–1085 16. Krutzfeldt A, Spahr R, Mertens S, Siegmund B, Piper HM (1990) Metabolism of exogenous substrates by coronary endothelial cells in culture. J Mol Cell Cardiol 22(12):1393–1404 17. Teuwen LA, Geldhof V, Carmeliet P (2019) How glucose, glutamine and fatty acid metabolism shape blood and lymph vessel development. Dev Biol 447(1):90–102 18. Barankay T, Baumgartl H, Lubbers DW, Seidl E (1976) Oxygen pressure in small lymphatics. Pflugers Arch 366(1):53–59 19. De Bock K, Georgiadou M, Schoors S, Kuchnio A, Wong BW, Cantelmo AR et al (2013) Role of PFKFB3-driven glycolysis in vessel sprouting. Cell 154(3):651–663

Evaluation of Endothelial Bioenergetics 20. Zhang XE, Adderley SP, Breslin JW (2016) Activation of RhoA, but not Rac1, mediates early stages of S1P-induced endothelial barrier enhancement. PLoS One 11(5):e0155490 21. Caja S, Enriquez JA (2017) Mitochondria in endothelial cells: sensors and integrators of environmental cues. Redox Biol 12:821–827 22. Tang X, Luo YX, Chen HZ, Liu DP (2014) Mitochondria, endothelial cell function, and vascular diseases. Front Physiol 5:175 23. Widlansky ME, Gutterman DD (2011) Regulation of endothelial function by mitochondrial reactive oxygen species. Antioxid Redox Signal 15(6):1517–1530 24. Dedkova EN, Ji X, Lipsius SL, Blatter LA (2004) Mitochondrial calcium uptake stimulates nitric oxide production in mitochondria of bovine vascular endothelial cells. Am J Physiol Cell Physiol 286(2):C406–C415 25. Tyagi N, Ovechkin AV, Lominadze D, Moshal KS, Tyagi SC (2006) Mitochondrial mechanism of microvascular endothelial cells apoptosis in hyperhomocysteinemia. J Cell Biochem 98(5):1150–1162 26. Madamanchi NR, Runge MS (2007) Mitochondrial dysfunction in atherosclerosis. Circ Res 100(4):460–473 27. Shenouda SM, Widlansky ME, Chen K, Xu G, Holbrook M, Tabit CE et al (2011) Altered mitochondrial dynamics contributes to endothelial dysfunction in diabetes mellitus. Circulation 124(4):444–453 28. Puddu P, Puddu GM, Cravero E, De Pascalis S, Muscari A (2007) The putative role of mitochondrial dysfunction in hypertension. Clin Exp Hypertens 29(7):427–434 29. Alves NG, Trujillo AN, Breslin JW, Yuan SY (2019) Sphingosine-1-phosphate reduces hemorrhagic shock and resuscitation-induced microvascular leakage by protecting endothelial mitochondrial integrity. Shock 52(4): 423–433 30. Sawant DA, Tharakan B, Hunter FA, Childs EW (2014) The role of intrinsic apoptotic signaling in hemorrhagic shock-induced microvascular endothelial cell barrier dysfunction. J Cardiovasc Transl Res 7(8):711–718 31. Yetkin-Arik B, Vogels IMC, Neyazi N, van Duinen V, Houtkooper RH, van Noorden CJF et al (2019) Endothelial tip cells in vitro are less glycolytic and have a more flexible response to metabolic stress than non-tip cells. Sci Rep 9(1):10414 32. Dranka BP, Hill BG, Darley-Usmar VM (2010) Mitochondrial reserve capacity in endothelial cells: the impact of nitric oxide and

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response to oxidative stress by metabolic profiling. Free Radic Biol Med 51(9): 1621–1635 49. Heinz S, Freyberger A, Lawrenz B, Schladt L, Schmuck G, Ellinger-Ziegelbauer H (2017) Mechanistic investigations of the mitochondrial complex I inhibitor rotenone in the context of pharmacological and safety evaluation. Sci Rep 7:45465 50. Hytti M, Korhonen E, Hyttinen JMT, Roehrich H, Kaarniranta K, Ferrington DA et al (2019) Antimycin A-induced mitochondrial damage causes human RPE cell death despite activation of autophagy. Oxidative Med Cell Longev 2019:1583656 51. Romero N, Swain P, Neilson A, Dranka BP (2017) Improving quantification of cellular glycolytic rate using Agilent Seahorse XF Technology. Agilent technologies White paper 52. Chaudhry R, Varacallo M (2020) Biochemistry, glycolysis. StatPearls, Treasure Island 53. Mookerjee SA, Goncalves RLS, Gerencser AA, Nicholls DG, Brand MD (2015) The contributions of respiration and glycolysis to extracellular acid production. Biochim Biophys Acta 1847(2):171–181 54. Pajak B, Siwiak E, Soltyka M, Priebe A, Zielinski R, Fokt I et al (2019) 2-Deoxy-dglucose and its analogs: from diagnostic to therapeutic agents. Int J Mol Sci 21(1):234 55. Romero N, Rogers G, Neilson A, Dranka BP (2018) Quantifying cellular ATP production rate using Agilent Seahorse XF Technology. Agilent technologies White paper

Chapter 21 Evaluation of Tight Junction Integrity in Brain Endothelial Cells Using Confocal Microscopy Himakarnika Alluri, Chander Sekhar Peddaboina, and Binu Tharakan Abstract The blood-brain barrier (BBB) is a highly complex and dynamic microvascular barrier that protects the brain parenchyma from the entry of pathogens, toxins, and other macromolecules and is a critical structure that helps to maintain brain homeostasis. The BBB is formed mainly by brain capillary endothelial cells and perivascular astrocytes and pericytes. One of the primary properties of the BBB is a tight regulation of paracellular permeability due to the presence of tight junctional complexes (also, adherens and gap junctions) between the neighboring microvascular endothelial cells. Alterations in the assembly of the tight junctions impair BBB properties, particularly influenced barrier integrity and permeability. The tight junctions of the BBB are mainly composed of proteins including claudins, occludin, and zonula occludens-1 (ZO-1). Zonula occludens-1 binds to the actin cytoskeleton, and its localization provides valuable information on the status of BBB integrity and permeability. Immunofluorescence localization of ZO-1 and/or other tight junction proteins is a reliable indicator of barrier integrity and permeability in microvascular endothelial cells. In microvascular endothelial cells, f-actin stress fiber formation significantly influences the rate and size of the inter-endothelial cell gap that form as cells retract from their borders. Rhodamine phalloidin is a popular conjugate used as a fluorescent label for f-actin. Herein, we describe the procedures for ZO-1 immunofluorescence and f-actin labeling followed by confocal microscopic imaging to determine barrier integrity and tight junction organization in brain microvascular endothelial cells in vitro. Key words Tight junction proteins, Blood-brain barrier, Zonula occludens-1 immunofluorescence, f-actin

1

Introduction The blood-brain barrier (BBB), predominantly formed by cerebrovascular endothelial cells and other cellular components of the neurovascular unit, creates a boundary between the bloodstream and brain parenchyma and provides protection to the brain from traumatic, ischemic, and other pathological conditions [1–4]. The BBB forms one of the tightest barriers in the human body blocking the transport of nearly all macromolecular and small molecular drugs into the brain [5]. Studies by Paul Ehlrich and Edwin Goldman in the late nineteenth and early twentieth century on barrier

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function in the brain paved way for our current understanding of the BBB a complex interaction of the cerebral microvascular endothelial cells and the tight junctions between them [6–8]. Endothelial cells restrict endocytosis and transcytosis within the brain acting as “transport barrier,” and they also metabolize many toxic and neuroactive substances, thus functioning as “metabolic barrier” [9]. Therapeutic molecules cannot transport across to the brain due to the tightness of BBB, and efforts are being made to permit their transportation by transiently opening the barrier using various methods [10]. Tight junction proteins, the structural and functional units of BBB, are expressed at a very early stage of embryonic development, and they undergo several posttranslational modifications (PTMs) to regulate endothelial cell functions like polarity, gene expression, signal transduction, and differentiation. Tight junction proteins are composed of various bicellular and tricellular proteins including claudins, occludin, zonula occludens, and tricellulins [11– 13]. Some of the cytoplasmic proteins like cingulin, afadin (AF6), 7H6, and zonula occludens (ZO)-1, (ZO)-2, and (ZO)-3 which bind to the actin cytoskeleton with the cell are part of the tight junction family proteins [14–16]. Zonula occludens (ZO) are multidomain scaffolding cytoplasmic phosphoproteins that belong to membrane-associated guanylate kinases (MAGUK) family, and they link the transmembrane tight junction proteins like claudin to the actin cytoskeleton via actin-binding proteins cingulin and afadin. Zonula occludens are expressed as ZO-1, ZO-2, or ZO-3 isoforms sharing sequence homology, and their C terminal domains are important for tight junction localization and interaction with other tight junction proteins and immunoglobulins [16, 17]. ZO proteins regulate tight junction integrity, cell-cell interaction and adhesion, and signal transduction, and based on the recent studies they also play a major role in the regulation of vascular permeability within the BBB [18, 19]. Brain trauma, peripheral vascular disease, and myocardial infarction lead to inflammation and edema resulting in loss of homeostatic balance and free radical generation causing alterations to blood-brain barrier tight junctions and microvascular permeability [20]. The current immunofluorescence technique will help assess the tight junction (TJ) integrity and stress fiber formation in endothelial cells. Immunofluorescence staining for ZO-1 protein is performed to determine TJ integrity and rhodamine phalloidin labeling for f-actin stress fiber formation in endothelial cells.

Confocal Microscopic Imaging of Endothelial Tight Junctions

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Materials 1. Rat brain microvascular endothelial cells (RBMVECs) [Cell Applications Inc., San Diego, CA]. 2. RBMVEC media [Cell Applications Inc., San Diego, CA]. 3. Nunc Lab Tek II-CC, 8-well glass chamber slides [Sigma Aldrich, St. Louis, MO]. 4. 15 mL polypropylene conical centrifuge tubes [Corning, NY]. 5. Fibronectin [Sigma Aldrich, St. Louis, MO]. 6. Bovine serum albumin [Sigma Aldrich, St. Louis, MO]. 7. Rhodamine phalloidin Carlsbad, CA].

[Thermo

Fisher

Scientific,

8. Rabbit anti ZO-1 [Thermo Fisher Scientific, Carlsbad, CA]. 9. Opti-MEM (1×)/reduced serum medium [Thermo Fisher Scientific, Carlsbad, CA]. 10. HyClone Dulbecco’s phosphate-buffered saline (PBS, without calcium, magnesium, or Phenol Red) [Thermo Fisher Scientific, Carlsbad, CA]. 11. Anti-rabbit IgG-FITC secondary antibody [Santa Cruz Biotechnology, Inc. Santa Cruz, CA]. 12. Vector VECTASHIELD® Mounting Media with DAPI [Vector Laboratories, Burlingame, CA].

3

Methods

3.1 Endothelial Cell Seeding

1. Culture RBMVEC in RBMVEC growth media on a 10 cm Petri dish coated with 5% fibronectin (50 μg/ml in PBS), replenish the media every 2 days until 80% confluency. 2. Once the cells reach desired confluency, wash the cell layer with 10 mL phosphate-buffered saline (PBS) twice and add 1 mL of 0.25% trypsin-ethylenediaminetetraacetic acid (EDTA) solution equilibrated to 37 °C. 3. Incubate at 37 °C in a tissue culture incubator for 2–5 min until the cells detach from the Petri dish. 4. Add 9 mL of complete media containing fetal bovine serum to neutralize the enzymatic activity of trypsin. Using a 10 mL serological pipette, gently collect the cell suspension in a 15 mL centrifuge tube and centrifuge at 1200 rpm for 5 min. 5. Aspirate the supernatant and resuspend the cell pellet at 50,000 cells/mL in fresh rat brain endothelial cell growth media.

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6. Transfer 300 μL of cell suspension into 5% fibronectin precoated Nunc Lab Tek II-CC 8-well sterile, glass chamber slides with 0.7 cm2/well surface area. 7. Grow the cells at 37 °C until confluence is achieved. 3.2 ZO-1 Immunofluorescence and Rhodamine Phalloidin Labeling for f-actin

1. Chamber slides containing RBMVEC monolayers are exposed to Opti-MEM/reduced serum media for an hour. 2. Wash the slides with 0.5 mL of phosphate-buffered saline (PBS) for a total of three washes. 3. Fix the cells using 4% paraformaldehyde in PBS for 15 min, followed by three washes in PBS. 4. Cells are permeabilized using 0.5% Triton X-100 in PBS for another 15 min. Block with 2% bovine serum albumin (BSA) in PBS for 1 h at room temperature. 5. After this step cells are either stained for ZO-1 or f-actin. 6. For ZO-1 immunofluorescence, staining cells are incubated with 100 μL of anti-rabbit primary antibody against ZO-1 diluted at 1:150 prepared in 2% BSA-PBS, overnight at 4 °C. 7. The following day, cells are washed in PBS with 1-min incubation for a total of three washes followed by incubation with fluorescein isothiocyanate (FITC)-tagged anti-rabbit secondary antibody diluted at 1:100 in 2% BSA-PBS for 1 h at room temperature. 8. For rhodamine phalloidin staining: Following blocking, cells are exposed to rhodamine phalloidin in 1:50 dilution, prepared in 2% BSA-PBS, for 20 min. 9. Cells from immunofluorescence and rhodamine phalloidin staining are washed with 0.5 mL PBS for a total of four washes and mounted using Vectashield antifade reagent containing DAPI. 10. Cells are visualized using a 60× water immersion lens and single optical slices on a confocal microscope (Fig. 1).

4

Notes 1. Precoat the culture dishes, chamber slides with fibronectin at 50 μg/mL in PBS for 1 h at 37 °C before seeding the RBMVEC. 2. Do not expose RBMVEC to 0.25% trypsin-EDTA solution for longer than 5 min, if the cells do not detach within 5 min, gently tap the culture plates to dislodge the cells from the surface.

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ZO-1

A

f-actin

B

Control

C

IL-1β

Calpain inhibitor III + IL-1β

Calpain inhibitor III

D

Fig. 1 Immunofluorescence localization of ZO-1 (a) and rhodamine phalloidin labeling of f-actin (b) in rat brain microvascular endothelial cells. ZO-1 tight junctional integrity and F-actin stress fiber formation were assessed using immunofluorescence localization and rhodamine phalloidin techniques, respectively. Exposure of the cells to interleukin 1β (IL-1β)-induced ZO-1 junctional disruption and F-actin stress fiber formation. The effect was reduced by pretreatment with calpain inhibitor III (a–d). IL-1β treatment-induced ZO-1 junctional disruption and F-actin stress fiber formation are shown by white arrows in A and B, respectively. The changes in ZO-1 localization and the formation of F-actin stress fibers were further determined using ImageJ (C and D, respectively). (The data are taken from a previously published study from our lab (Alluri et al. [21]) Creative Commons License https://creativecommons.org/licenses/)

3. Once the slides are mounted with Vectashield, seal the slides with nail polish to avoid the slides from drying.

Acknowledgments The authors acknowledge the support from the National Institutes of Health (NIH) grant 5 SC3 NS127765-02 (BT).

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References 1. Dietrich W, Erbguth F (2013) Increased intracranial pressure and brain edema. Med Klin Intensivmed Notfmed 108(2):157–169; quiz 170–171 2. Bundgaard M, Abbott NJ (2008) All vertebrates started out with a glial blood-brain barrier 4-500 million years ago. Glia 56(7): 699–708 3. Cecchelli R et al (2007) Modelling of the blood-brain barrier in drug discovery and development. Nat Rev Drug Discov 6(8): 650–661 4. Guest J et al (2013) Relationship between central and peripheral fatty acids in humans. Lipids Health Dis 12:79 5. Pardridge WM (2005) The blood-brain barrier: bottleneck in brain drug development. NeuroRx 2(1):3–14 6. Mayhan WG (2001) Regulation of blood-brain barrier permeability. Microcirculation 8(2): 89–104 7. Nag S, Kapadia A, Stewart DJ (2011) Review: molecular pathogenesis of blood-brain barrier breakdown in acute brain injury. Neuropathol Appl Neurobiol 37(1):3–23 8. Engelhardt B, Sorokin L (2009) The bloodbrain and the blood-cerebrospinal fluid barriers: function and dysfunction. Semin Immunopathol 31(4):497–511 9. Abbott NJ, Ronnback L, Hansson E (2006) Astrocyte-endothelial interactions at the blood-brain barrier. Nat Rev Neurosci 7(1): 41–53 10. Stamatovic SM, Keep RF, Andjelkovic AV (2008) Brain endothelial cell-cell junctions: how to “open” the blood brain barrier. Curr Neuropharmacol 6(3):179–192 11. Daneman R et al (2010) The mouse bloodbrain barrier transcriptome: a new resource for understanding the development and

function of brain endothelial cells. PLoS One 5(10):e13741 12. Annunziata P et al (2002) Substance P antagonist blocks leakage and reduces activation of cytokine-stimulated rat brain endothelium. J Neuroimmunol 131(1–2):41–49 13. Mariano C et al (2013) Tricellulin expression in brain endothelial and neural cells. Cell Tissue Res 351(3):397–407 14. Paris L et al (2008) Structural organization of the tight junctions. Biochim Biophys Acta 1778(3):646–659 15. Thal SC et al (2012) Volatile anesthetics influence blood-brain barrier integrity by modulation of tight junction protein expression in traumatic brain injury. PLoS One 7(12): e50752 16. Fanning AS et al (1998) The tight junction protein ZO-1 establishes a link between the transmembrane protein occludin and the actin cytoskeleton. J Biol Chem 273(45): 29745–29753 17. Bazzoni G, Dejana E (2004) Endothelial cellto-cell junctions: molecular organization and role in vascular homeostasis. Physiol Rev 84(3):869–901 18. Gonzalez-Mariscal L, Betanzos A, Avila-Flores A (2000) MAGUK proteins: structure and role in the tight junction. Semin Cell Dev Biol 11(4):315–324 19. Arshad F et al (2010) Blood-brain barrier integrity and breast cancer metastasis to the brain. Pathol Res Int 2011:920509 20. Kaur C, Ling EA (2008) Blood brain barrier in hypoxic-ischemic conditions. Curr Neurovasc Res 5(1):71–81 21. Alluri H, Grimsley M, Shaji CA et al (2016) Attenuation of blood-brain barrier breakdown and hyperpermeability by calpain inhibition. J Biol Chem 291:26958–26969

INDEX A Actin cytoskeleton....................................... 187, 236, 258 Actin stress fibers......................................... 131, 138, 141 Adhesion ............................. 3, 48–50, 54, 56, 57, 59, 78, 117, 164, 170, 177, 187, 225–232, 236, 258 Angiogenesis..........................63–74, 105–115, 148, 149, 185, 186, 189, 199, 210, 243 Aortic rings................................... 64, 106, 107, 112, 114 ATP rate................................................................ 243, 251

Endothelium................................ 1–3, 13, 34, 36, 78, 89, 100, 102, 147, 148, 163–165, 185, 199, 225–232, 241–243 Extravasation ....................... 45, 120, 177–184, 225, 235

F FITC dextran permeability assay.................................. 206

G Glycocalyx............................................................. 163–173 Glycolysis .............................................................. 241–254

B Barrier function ...........................35, 47, 53, 54, 60, 189, 242, 243, 257 Blood-brain barrier (BBB)......................... 59, 60, 77–87, 117–126, 177–184, 199–203, 235, 236, 257, 258 BSI-lectin ........................................................73, 165, 169

C Cannulation .......................21–26, 29–31, 33–36, 93–95, 165–167, 173 Capillary sprouting............................................. 64, 70, 72 Chorioallantoic membrane ............................64, 106, 188 COVID-19 endotheliitis ..................................... 185, 186 Cremaster muscle exposure surgery............................... 16

D Dextran .........................5, 14, 15, 17–19, 34, 79, 80, 84, 102, 118, 166, 170, 171, 206, 209, 210, 219 Donor eyes ........................................................... 206, 210 Dye-exclusion ...............................................165, 170–173

E Edema .........................1, 13, 39, 40, 118, 178, 185–187, 200, 206, 235, 258 Endothelial barrier ......................... 2, 5, 47–61, 130, 200 Endothelial barrier functions .................... 21, 40, 47, 48, 185–196, 242 Endothelial cells .............................. 1, 14, 40, 47, 63, 78, 106, 117, 129, 147, 163, 186, 199, 206, 225, 236, 242, 257 Endothelial surface layer...................................... 163–173

H Hemorrhagic shock (HS) ........................ 39, 40, 45, 164, 200, 242, 243 Human primary choroidal endothelial cells ....... 205–222 Human retinal pigment epithelial cells (hRPE) ................... 205–210, 213, 214, 216–220 Hypoxia ...............................................186–192, 194, 195

I Inflammation ...............................1, 13, 14, 90, 122, 147, 164, 185, 186, 189, 225, 226, 235, 258 Integrity .................................77–87, 118, 131, 164, 165, 167, 169, 177–179, 186, 192, 199, 200, 235–240, 243, 257–261 Intravital microscopy (IVM) ............... 39–41, 44, 89–91, 117–126, 164–167, 169–173, 188 In vitro blood retinal barrier model............................. 206 In vitro models .............................................................. 206 Ischemia.................................................... 63, 91, 96, 148, 186, 188, 189 Ischemia/reperfusion injury...................................89–102 Isolated vessel ............................................................31, 33

J Junctional-associated intermittent lamellipodia .......... 130

L Leakage ........................... 1–3, 13, 14, 17–19, 25, 34, 36, 40, 86, 118, 178, 179, 185, 191, 206

Binu Tharakan (ed.), Vascular Hyperpermeability: Methods and Protocols, Methods in Molecular Biology, vol. 2711, https://doi.org/10.1007/978-1-0716-3429-5, © Springer Science+Business Media, LLC, part of Springer Nature 2024

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Leukocyte-endothelial interaction .........................89–102 Local lamellipodia ...............................130, 131, 138, 139

M Matrigel ...................................... 105–115, 149, 155, 158 Mesentery ...................................... 22, 27, 28, 43, 44, 46, 63–74, 122, 164, 165, 169, 173 Metabolism............................................79, 241, 243, 244 Microcirculation .............................. 3, 91, 102, 123, 124, 163–165, 171 Microfluidics....................................................... 64, 77–87 Mitochondria........................................................ 242, 243

O Organ-on-chip ................................................................ 79

P Perfusion......................................3, 32, 40, 79, 130, 131, 134, 138, 152, 182, 183, 200 Permeability..................................... 1, 13, 21, 39, 47, 78, 102, 118, 129, 164, 178, 185, 199, 200, 206, 236, 258

Pressure..................................2, 3, 21, 25, 27, 30–35, 41, 43, 44, 72, 92, 93, 95, 99, 118, 121, 126, 152, 167, 168, 178, 188, 200, 207

S Seahorse ................................................................ 241–254 Sepsis................................................. 1, 47, 185, 186, 225

T Time-lapse imaging.................... 64, 68–70, 72, 130, 189 Tissue culture .......................................66, 207, 213, 214, 226, 228, 229, 259 Transwell inserts...................................... 4, 6–8, 201, 203 Tumor inflammation..................................................... 225 Tumor metastasis .......................................................... 225 Two-dimensional angiogenesis .................. 106, 107, 110 Two-photon intravital imaging ................................18, 93 Two-photon intravital microscopy................................. 93

V Vascular permeability ..................................13–19, 21–37, 39–41, 102, 120, 125, 179, 185–189, 193, 199, 258