Cystic Fibrosis: Diagnosis and Protocols, Volume II: Methods and Resources to Understand Cystic Fibrosis (Methods in Molecular Biology 742)

ME T H O D S IN MO L E C U L A R BI O L O G Y Series Editor John M. Walker School of Life Sciences University of Her...

Author: Margarida D. Amaral | Karl Kunzelmann

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ME T H O D S

IN

MO L E C U L A R BI O L O G Y

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

For other titles published in this series, go to www.springer.com/series/7651

TM

Cystic Fibrosis Diagnosis and Protocols, Volume II: Methods and Resources to Understand Cystic Fibrosis

Edited by

Margarida D. Amaral Centre for Biodiversity and Functional and Integrative Genomics, Faculty of Sciences, University of Lisboa, Lisboa, Portugal

Karl Kunzelmann Department of Physiology, University of Regensburg, Regensburg, Germany

Editors Margarida D. Amaral Centre for Biodiversity & Functional and Integrative Genomics University of Lisboa Lisboa 1749-016, Portugal [email protected]

Karl Kunzelmann Department of Physiology University of Regensburg Regensburg 93053, Germany [email protected]

ISSN 1064-3745 e-ISSN 1940-6029 ISBN 978-1-61779-119-2 e-ISBN 978-1-61779-120-8 DOI 10.1007/978-1-61779-120-8 Springer New York Dordrecht Heidelberg London Library of Congress Control Number: 2011925926 © Springer Science+Business Media, LLC 2011 All rights reserved. This work may not be translated or copied in whole or in part without the written permission of the publisher (Humana Press, c/o Springer Science+Business Media, LLC, 233 Spring Street, New York, NY 10013, USA), except for brief excerpts in connection with reviews or scholarly analysis. Use in connection with any form of information storage and retrieval, electronic adaptation, computer software, or by similar or dissimilar methodology now known or hereafter developed is forbidden. The use in this publication of trade names, trademarks, service marks, and similar terms, even if they are not identified as such, is not to be taken as an expression of opinion as to whether or not they are subject to proprietary rights. While the advice and information in this book are believed to be true and accurate at the date of going to press, neither the authors nor the editors nor the publisher can accept any legal responsibility for any errors or omissions that may be made. The publisher makes no warranty, express or implied, with respect to the material contained herein. Printed on acid-free paper Humana Press is part of Springer Science+Business Media (www.springer.com)

Foreword This book represents a milestone in the worldwide cystic fibrosis (CF) community’s efforts to continue to pave the way toward the development of new and innovative therapies that address the basic defect in CF. But no book on this subject would be possible without the invaluable contributions of the many patients, families, and disease-related organizations that played a key role in creating the science outlined in these chapters. As an orphan disease, CF does not receive sufficient funding from traditional supporting agencies but depends instead on a vast network of people who selflessly give their time and energy to raise the dollars to support the research that will lead to new treatments and a cure. Much of the science described in the pages that follow is the result of funds raised by the CF community, as well as the willingness of patients to provide tissue specimens, share their data in patient registries, and participate in clinical studies. These contributions have been critical to the success that CF research is experiencing worldwide. The global cystic fibrosis community is clearly unique and has often been described as a “culture of research.” Shared among the many patient groups that represent about 70,000 people with CF worldwide is the belief that we will ultimately cure this disease through research. The constancy of this shared mission to cure CF by focusing on research is part of what sets the CF community apart from other rare diseases. The clear promise of small molecules and the excitement over gene therapy have kindled a sense of optimism that is critical to sustaining the momentum toward finding a cure. In addition to the consistent focus on research, there are a number of other unique traits that the CF community around the world possesses that make it one of a kind. Some of the distinguishing qualities include the following: • Willingness to share: Because of the insidious nature of CF, there is a rare sense of cooperative spirit among scientists, physicians, caregivers, patients, and families all over the world who are dedicated to a cure. CF research data know no borders, and waiting until data are published is not part of the CF research culture. The advancement of science is an open book, and the rapid exchange of new ideas and approaches is a mainstay of CF conferences and workshops in North America and overseas. • Willingness to take risks: The search for the gene in the 1980s is an example of the risks and rewards of the pioneering work of the global CF populace. In the early 1980s, CF communities began to collect blood and tissue samples from families with multiplyaffected individuals with CF. With newly evolved technologies (such as chromosome jumping), which could be quickly applied to these samples, and the rapid exchange of information, numerous efforts to find the gene were launched. The discovery of the CF gene in 1989 was the result of an intricate and highly successful international collaboration and is hailed today as one of the major milestones in modern medical research. Because of the involvement of CF families, patients, and the US Cystic Fibrosis Foundation, this groundbreaking discovery occurred over 14 years prior to the publication of the human genome. Importantly, it gave scientists an opportunity to explore the relationship of the genetic defect with the pathogenesis associated with CF. This discovery was a prelude to the effort to move from a knowledge acquisition

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Foreword mode of research to the current CF research efforts, whereby scientists are using acquired knowledge about CF to develop new approaches to treat the disease.

• Willingness to take responsibility for its own destiny: Because there are only 70,000 CF patients throughout the world, there has been reluctance in the biopharmaceutical arena to enter the field of CF. Without their involvement, the ability to develop novel therapeutics is limited. In the late 1990s, frustrated by this fact, the US CF Foundation dramatically changed its business model and created a program to reduce risk of industry involvement in CF research by providing early funding and access to scientific and clinical expertise. This successful array of alliances with industry has led to an exciting clinical pipeline of products, including small molecules that are now being tested in centers worldwide, any of which could have a profound impact on individuals with CF. Similarly, upon realizing industry’s dwindling interest in gene therapy for CF, the British Trust launched a Gene Therapy Consortium that has painstakingly worked through some of the critical issues and problems associated with applying this pioneering mode of therapy for CF. As a result, and with a significant financial investment by the British Trust, the most comprehensive gene therapy clinical trial process is underway in the British Isles. More recently, clinical trial networks have been established throughout North America and Europe to facilitate the evaluation of new clinical entities in order to hasten the regulatory process leading to drug approval. These clinical trial networks are linked through the sharing of data, expertise, and experience to contribute to the worldwide clinical trial efforts. These are just a few examples of the willingness of the CF community to make strategic investments, some of which, in the case of other diseases, would be taken by industry or the government to bring us closer to accomplishing our mission. • Willingness to accept responsibility for the coordinating role in the areas of care, teaching, and research: In many countries, scientists and clinicians look to the government for funding – agencies like the US National Institutes of Health and other equivalent organizations. However, not only is the science of CF frequently funded by CF organizations, but its direction is often defined by these entities as well. Similarly, the outstanding care that is provided all over the world is driven by the standards and guidelines set by these universal CF organizations. These guidelines are commonly established in international forums sponsored by CF groups. Once again, the community looks to CF organizations for leadership. These volumes are the result of a distinct and worldwide undertaking. The environment, funding, and culture that have been put in place by patient organizations, coupled with the ability to bring the best minds to the field of CF, have made the science described in this book possible. This publication will be a useful tool as we continue to translate the knowledge acquired during decades of basic research to the development of new therapies that will modify and change the course of the disease in CF patients in the years ahead. Cystic Fibrosis: Diagnosis and Protocols is the fulfillment of decades of hard work by the volunteers and staff of the patient groups and organizations that have helped to pave the way toward our ultimate goal: a cure and control for cystic fibrosis. Bethesda, MD, USA

Robert J. Beall

Preface More than 20 years have passed since the identification of the gene responsible for cystic fibrosis (CF) and undoubtedly many milestones have highlighted this area of research. But we have to admit it, progress towards finding a way of curing the disease has been slower than we initially expected and wished. Apparently, this is not due to a lack of research efforts in the field, since in recent years, the CF research community has been producing on average ∼1,500 papers annually. So, probably we still need to dig deeper and with better tools to understand further the basic biological mechanisms underlying this complex disease. Nevertheless, it is increasingly difficult to grasp and use the already wide and still growing range of diverse methods currently employed to study CF so as to understand it in its multidisciplinary nature. The aim of these Cystic Fibrosis: Diagnosis and Protocols volumes is thus to provide the CF research community (and that in related fields) with a very wide range of high-quality experimental tools, as an easy way to grasp and use classical and novel methods applied to CF. Hence, it is expected that it will contribute to accelerate the advancement of knowledge in this area. The purpose is thus to offer selected “good practice protocols” with a level of technical detail which is rarely published in peer-reviewed journals. Moreover, it is expected that this information will also enable researchers to identify subtle differences regarding techniques in their own laboratories, which often account for apparently “contradictory” data in the literature. Co-authorship from both sides of the Atlantic was particularly encouraged. In the 2002 edition of this volume and in another previous comprehensive compilation of Methods for Cystic Fibrosis and CFTR Research,1 a large set of classical techniques used for CF research were already covered. So, here the focus is placed on innovative methodologies (some revolutionizing our way of doing science) by describing in detail how to perform and exploit these emergent techniques applied to CF. Moreover, a complete section has been devoted to available resources such as useful software and databases, as well as cell lines and animal models, reviewed for their usefulness towards multiple purposes. Notwithstanding, the more “classical” methods can also undergo improvements and thus their most up-to-date and revised versions are also recapped here by the leading experts. All book sections are introduced by an overview discussing the applicability and practicality of the protocols with examples. It is hoped that the methods presented and revised here will provide users with optimal working tools to address their pressing questions in the best technical way while helping

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Journal of Cystic Fibrosis (2004), volume 3 (Supplement 2), a special issue focused on “Methods for Cystic Fibrosis and CFTR Research” and The online “Virtual Repository of the Cystic Fibrosis European Network” at: http://central.igc.gulbenkian.pt/cftr/vr/index.htm

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all of us, as a research and clinical community, to move faster hand-in-hand towards unravelling the secrets of this (and possibly other) challenging disorder(s) and cure it. Finally, we wish to thank all authors for their enthusiasm in joining us in this project by contributing with their best protocols to this book and also for their patience with our multiple requests. Special thanks to Renata Vincent for her help in dealing with the manuscripts. Moreover, we would like to express our gratitude to the whole CF community in general, researchers, clinicians and all caregivers and other professionals, not forgetting CF patients and their families, for their continuous efforts towards finding a way out of this still devastating disease. We believe that we will be there soon and we hope this book somehow contributes to getting there sooner. Then, when our goals are met, all efforts will have been worthwhile, or as the Portuguese poet Fernando Pessoa has put it, “All is worthwhile if the soul is not small”. Lisboa, Portugal Regensburg, Germany

Margarida D. Amaral Karl Kunzelman

Contents Foreword . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

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Preface . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

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Contributors . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

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SECTION I: 1.

PATHOPHYSIOLOGY OF CYSTIC FIBROSIS

Introduction to Section I: Overview of Approaches to Study Cystic Fibrosis Pathophysiology . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Mark T. Clunes and Richard C. Boucher

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Imaging CFTR Protein Localization in Cultured Cells and Tissues . . . . . . . . Silvia M. Kreda and Martina Gentzsch

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CFTR Regulation of Epithelial Sodium Channel . . . . . . . . . . . . . . . . . . Yawar J. Qadri, Estelle Cormet-Boyaka, Dale J. Benos, and Bakhrom K. Berdiev

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Methods for Evaluating Inflammation in Cystic Fibrosis . . . . . . . . . . . . . . Assem G. Ziady and Pamela B. Davis

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Methods for ASL Measurements and Mucus Transport Rates in Cell Cultures . . . Erin N. Worthington and Robert Tarran

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Measurement of Fluid Secretion from Intact Airway Submucosal Glands . . . . . Jeffrey J. Wine, Nam Soo Joo, Jae Young Choi, Hyung-Ju Cho, Mauri E. Krouse, Jin V. Wu, Monal Khansaheb, Toshiya Irokawa, Juan Ianowski, John W. Hanrahan, Alan W. Cuthbert, and Kim V. Tran

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Measurements of Intracellular Calcium Signals in Polarized Primary Cultures of Normal and Cystic Fibrosis Human Airway Epithelia . . . . . . . . . 113 Carla M.P. Ribeiro

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Identification and Quantification of Mucin Expression . . . . . . . . . . . . . . . 127 Kristina A. Thomsson and Gunnar C. Hansson

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Methods to Classify Bacterial Pathogens in Cystic Fibrosis . . . . . . . . . . . . . 143 Thomas Bjarnsholt, Xiaohui Chen Nielsen, Ulla Johansen, Lena Nørgaard, and Niels Høiby

10. Approaches to Study Differentiation and Repair of Human Airway Epithelial Cells . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 173 Sophie Crespin, Marc Bacchetta, Song Huang, Tecla Dudez, Ludovic Wiszniewski, and Marc Chanson

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SECTION II: OMIC APPROACHES TO STUDY CYSTIC FIBROSIS 11. Introduction to Section II: Omics in the Biology of Cystic Fibrosis . . . . . . . . 189 William E. Balch 12. Microarray mRNA Expression Profiling to Study Cystic Fibrosis . . . . . . . . . . 193 Shyam Ramachandran, Luka A. Clarke, Todd E. Scheetz, Margarida D. Amaral, and Paul B. McCray Jr. 13. Quantitative Differential Proteomics of Cystic Fibrosis Cell Models by SILAC (Stable Isotope Labelling in Cell Culture) . . . . . . . . . . . . . . . . 213 Ida Chiara Guerrera, Mario Ollero, Diane-Lore Vieu, and Aleksander Edelman 14. Application of Mass Spectrometry to Study Proteomics and Interactomics in Cystic Fibrosis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 227 William E. Balch and John R. Yates III 15. Functional Genomics Assays to Study CFTR Traffic and ENaC Function . . . . . 249 Joana Almaça, Shehrazade Dahimène, Nicole Appel, Christian Conrad, Karl Kunzelmann, Rainer Pepperkok, and Margarida D. Amaral 16. New Lipidomic Approaches in Cystic Fibrosis . . . . . . . . . . . . . . . . . . . 265 Mario Ollero, Ida Chiara Guerrera, Giuseppe Astarita, Daniele Piomelli, and Aleksander Edelman SECTION III: RESOURCES 17. Introduction to Section III: Resources for CFTR Research . . . . . . . . . . . . 281 Margarida D. Amaral 18. Primary Epithelial Cell Models for Cystic Fibrosis Research . . . . . . . . . . . . 285 Scott H. Randell, M. Leslie Fulcher, Wanda O’Neal, and John C. Olsen 19. Comparative Biology of Cystic Fibrosis Animal Models . . . . . . . . . . . . . . 311 John T. Fisher, Yulong Zhang, and John F. Engelhardt 20. CFTR Folding Consortium: Methods Available for Studies of CFTR Folding and Correction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 335 Kathryn W. Peters, Tsukasa Okiyoneda, William E. Balch, Ineke Braakman, Jeffrey L. Brodsky, William B. Guggino, Christopher M. Penland, Harvey B. Pollard, Eric J. Sorscher, William R. Skach, Philip J. Thomas, Gergely L. Lukacs, and Raymond A. Frizzell 21. Evaluation of the Disease Liability of CFTR Variants . . . . . . . . . . . . . . . . 355 Patrick R. Sosnay, Carlo Castellani, Mary Corey, Ruslan Dorfman, Julian Zielenski, Rachel Karchin, Christopher M. Penland, and Garry R. Cutting Index . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 373

Contributors JOANA ALMAÇA • Faculty of Sciences, BioFiG-Centre for Biodiversity and Functional and Integrative Genomics, University of Lisboa, Lisboa, Portugal; Department of Genetics, Centre of Human Genetics, National Institute of Health, Lisboa, Portugal MARGARIDA D. AMARAL • Faculty of Sciences, BioFiG-Centre for Biodiversity, Functional and Integrative Genomics, University of Lisboa, Lisboa, Portugal; Department of Genetics, Centre of Human Genetics, National Institute of Health, Lisboa, Portugal; EMBL-European Molecular Biology Laboratory, Heidelberg, Germany NICOLE APPEL • EMBL-European Molecular Biology Laboratory, Heidelberg, Germany GIUSEPPE ASTARITA • Drug Discovery and Development Unit, Italian Institute of Technology, Genoa, Italy; Department of Pharmacology, University of California, Irvine, CA, USA MARC BACCHETTA • Laboratory of Clinical Investigation III, Faculty of Medicine, Department of Pediatrics, Geneva University Hospitals and University of Geneva, Foundation for Medical Research, Geneva, Switzerland WILLIAM E. BALCH • Departments of Cell Biology, Molecular Biology and Chemical Physiology, The Skaggs Institute for Chemical Biology and the Institute for Childhood and Neglected Disease, The Scripps Research Institute, La Jolla, CA, USA; The Institute for Childhood and Neglected Disease, The Scripps Research Institute, La Jolla, CA, USA DALE J. BENOS • Physiology and Biophysics, University of Alabama at Birmingham, Birmingham, AL, USA BAKHROM K. BERDIEV • Department of Cell Biology and Gregory Fleming James Cystic Fibrosis Research Center, University of Alabama at Birmingham, Birmingham, AL, USA THOMAS BJARNSHOLT • Department of Clinical Microbiology, University of Copenhagen, Copenhagen, DenmarkDanish CF Centre, Rigshospitalet, University of Copenhagen, Copenhagen, Denmark RICHARD C. BOUCHER • Cystic Fibrosis/Pulmonary Research and Treatment Center, University of North Carolina at Chapel Hill, Chapel Hill, NC, USA INEKE BRAAKMAN • Cellular Protein Chemistry, Utrecht University, Utrecht, The Netherlands JEFFREY L. BRODSKY • Department of Biological Sciences, University of Pittsburgh, Pittsburgh, PA, USA CARLO CASTELLANI • Cystic Fibrosis Centre, Ospedale Civile Maggiore, Verona, Italy MARC CHANSON • Laboratory of Clinical Investigation III, Faculty of Medicine, Department of Pediatrics, Geneva University Hospitals and University of Geneva, Foundation for Medical Research, Geneva, Switzerland HYUNG-JU CHO • Cystic Fibrosis Research Laboratory, Stanford University, Stanford, CA, USA JAE YOUNG CHOI • Cystic Fibrosis Research Laboratory, Stanford University, Stanford, CA, USA; Department of Otorhinolaryngology, Yonsei University, Seoul, Korea

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LUKA A. CLARKE • Faculty of Sciences, BioFIG-Centre for Biodiversity, Functional and Integrative Genomics, University of Lisboa, Lisbon, Portugal MARK T. CLUNES • Department of Physiology and Neuroscience, St. George’s University, True Blue Campus, Grenada, West Indies CHRISTIAN CONRAD • EMBL-European Molecular Biology Laboratory, Heidelberg, Germany MARY COREY • The Hospital for Sick Children, University of Toronto, Toronto, ON, Canada ESTELLE CORMET-BOYAKA • Division of Pulmonary, Critical Care, and Sleep Medicine, Davis Heart and Lung Research Institute, The Ohio State University, Columbus, OH, USA SOPHIE CRESPIN • Laboratory of Clinical Investigation III, Faculty of Medicine, Department of Pediatrics, Geneva University Hospitals, and University of Geneva, Foundation for Medical Research, Geneva, Switzerland ALAN W. CUTHBERT • Department of Medicine, University of Cambridge, Addenbrooke’s Hospital, Cambridge, UK GARRY R. CUTTING • McKusick-Nathans Institute of Genetic Medicine, Johns Hopkins University School of Medicine, Baltimore, MD, USA SHEHRAZADE DAHIMÈNE • Faculty of Sciences, BioFiG-Centre for Biodiversity and Functional and Integrative Genomics, University of Lisboa, Lisboa, Portugal PAMELA B. DAVIS • Department of Pediatrics, Case Western Reserve University, Cleveland, OH, USA RUSLAN DORFMAN • The Hospital for Sick Children, University of Toronto, Toronto, ON, Canada TECLA DUDEZ • Laboratory of Clinical Investigation III, Faculty of Medicine, Department of Pediatrics, Geneva University Hospitals and University of Geneva, Foundation for Medical Research, Geneva, Switzerland ALEKSANDER EDELMAN • INSERM U845, Université Paris Descartes, Paris, France JOHN F. ENGELHARDT • Department of Anatomy and Cell Biology, Carver College of Medicine, University of Iowa, Iowa City, IA, USA; Department of Internal Medicine, Carver College of Medicine, University of Iowa, Iowa City, IA, USA; Center for Gene Therapy, Carver College of Medicine, University of Iowa, Iowa City, IA, USA JOHN T. FISHER • Department of Anatomy and Cell Biology, Carver College of Medicine, University of Iowa, Iowa City, IA, USA RAYMOND A. FRIZZELL • Department of Cell Biology and Physiology, University of Pittsburgh School of Medicine, Pittsburgh, PA, USA M. LESLIE FULCHER • Cystic Fibrosis/Pulmonary Research and Treatment Center, The University of North Carolina at Chapel Hill, Chapel Hill, NC, USA MARTINA GENTZSCH • Cystic Fibrosis/Pulmonary Research and Treatment Center, University of North Carolina at Chapel Hill, Chapel Hill, NC, USA; Department of Cell and Developmental Biology, University of North Carolina at Chapel Hill, Chapel Hill, NC, USA IDA CHIARA GUERRERA • Plateau Protéomes, IFR94, Université Paris Descartes, Paris, France; INSERM U845, Université Paris Descartes, Paris, France WILLIAM B. GUGGINO • Department of Physiology, Johns Hopkins University, Baltimore, MD, USA JOHN W. HANRAHAN • Department of Physiology, McGill University, Montreal, QC, Canada

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GUNNAR C. HANSSON • Department of Medical Biochemistry, University of Gothenburg, Gothenburg, Sweden NIELS HØIBY • Department of Clinical Microbiology, University of Copenhagen, Copenhagen, Denmark; Danish CF Centre, Rigshospitalet, University of Copenhagen, Copenhagen, Denmark; Institute of International Health, Immunology and Microbiology, University of Copenhagen, Copenhagen, Denmark SONG HUANG • Epithelix, SàRL, Plan-les-Ouates, Switzerland JUAN IANOWSKI • Department of Physiology, University of Saskatchewan, Saskatoon, SK, Canada TOSHIYA IROKAWA • Health Administration Center, Tohoku University, Sendai, Japan ULLA JOHANSEN • Department of Clinical Microbiology, University of Copenhagen, Copenhagen, Denmark NAM SOO JOO • Cystic Fibrosis Research Laboratory, Stanford University, Stanford, CA, USA RACHEL KARCHIN • Department of Biomedical Engineering, Institute for Computational Medicine, Johns Hopkins University, Baltimore, MD, USA MONAL KHANSAHEB • Cystic Fibrosis Research Laboratory, Stanford University, Stanford, CA, USA SILVIA M. KREDA • Cystic Fibrosis/Pulmonary Research and Treatment Center, University of North Carolina at Chapel Hill, Chapel Hill, NC, USA MAURI E. KROUSE • Cystic Fibrosis Research Laboratory, Stanford University, Stanford, CA, USA KARL KUNZELMANN • Department of Physiology, University of Regensburg, Regensburg, Germany GERGELY L. LUKACS • Department of Physiology, McGill University, Montreal, QC, Canada PAUL B. MCCRAY JR • Interdisciplinary Program in Genetics, Department of Pediatrics, University of Iowa, Iowa City, IA, USA XIAOHUI CHEN NIELSEN • Department of Clinical Microbiology, University of Copenhagen, Copenhagen, Denmark LENA NØRGAARD • Department of Clinical Microbiology, University of Copenhagen, Copenhagen, Denmark TSUKASA OKIYONEDA • Department of Physiology, McGill University, Montreal, QC, Canada MARIO OLLERO • INSERM U845, Université Paris Descartes, Paris, France JOHN C. OLSEN • Cystic Fibrosis/Pulmonary Research and Treatment Center, The University of North Carolina at Chapel Hill, Chapel Hill, NC, USA WANDA O’NEAL • Cystic Fibrosis/Pulmonary Research and Treatment Center, The University of North Carolina at Chapel Hill, Chapel Hill, NC, USA CHRISTOPHER M. PENLAND • Cystic Fibrosis Foundation, Bethesda, MD, USA RAINER PEPPERKOK • EMBL-European Molecular Biology Laboratory, Heidelberg, Germany KATHRYN W. PETERS • Department of Cell Biology and Physiology, University of Pittsburgh, Pittsburgh, PA, USA DANIELE PIOMELLI • Drug Discovery and Development Unit, Italian Institute of Technology, Genoa, Italy; Department of Pharmacology, University of California, Irvine, CA, USA

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HARVEY B. POLLARD • Department of Anatomy, Physiology, and Genetics, Uniformed Services University of the Health Sciences, Bethesda, MD, USA YAWAR J. QADRI • Medical Scientist Training Program, University of Alabama at Birmingham, Birmingham, AL, USA SHYAM RAMACHANDRAN • Interdisciplinary Program in Genetics, Department of Pediatrics, University of Iowa, Iowa City, IA, USA SCOTT H. RANDELL • Cystic Fibrosis/Pulmonary Research and Treatment Center, The University of North Carolina at Chapel Hill, Chapel Hill, NC, USA CARLA M.P. RIBEIRO • Department of Medicine, Cystic Fibrosis/Pulmonary Research and Treatment Center, The University of North Carolina, Chapel Hill, NC, USA TODD E. SCHEETZ • Department of Ophthalmology and Visual Sciences, Interdisciplinary Program in Genetics, University of Iowa, Iowa City, IA, USA WILLIAM R. SKACH • Department of Biochemistry and Molecular Biology, Oregon Health & Science University, Portland, OR, USA ERIC J. SORSCHER • Gregory Fleming James Cystic Fibrosis Research Center, University of Alabama at Birmingham, Birmingham, AL, USA PATRICK R. SOSNAY • McKusick-Nathans Institute of Genetic Medicine, Johns Hopkins University School of Medicine, Baltimore, MD, USA ROBERT TARRAN • Cystic Fibrosis/Pulmonary Research and Treatment Center, The University of North Carolina, Chapel Hill, NC, USA PHILIP J. THOMAS • Department of Physiology, University of Texas Southwestern Medical Center, Dallas, TX, USA KRISTINA A. THOMSSON • Department of Medical Biochemistry, University of Gothenburg, Gothenburg, Sweden KIM V. TRAN • Cystic Fibrosis Research Laboratory, Stanford University, Stanford, CA, USA DIANE-LORE VIEU • INSERM U845, Université Paris Descartes, Paris, France JEFFREY J. WINE • Cystic Fibrosis Research Laboratory, Stanford University, Stanford, CA, USA LUDOVIC WISZNIEWSKI • Epithelix, SàRL, Plan-les-Ouates, Switzerland ERIN N. WORTHINGTON • Cystic Fibrosis/Pulmonary Research and Treatment Center, The University of North Carolina, Chapel Hill, NC, USA JIN V. WU • Cystic Fibrosis Research Laboratory, Stanford University, Stanford, CA, USA JOHN R. YATES III • Department of Chemical Physiology, The Scripps Research Institute, La Jolla, CA, USA YULONG ZHANG • Department of Anatomy and Cell Biology, Carver College of Medicine, University of Iowa, Iowa City, IA, USA ASSEM G. ZIADY • Department of Pediatrics, Case Western Reserve University, Cleveland, OH, USA JULIAN ZIELENSKI • The Hospital for Sick Children, University of Toronto, Toronto, ON, Canada

Section I Pathophysiology of Cystic Fibrosis

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Chapter 1 Introduction to Section I: Overview of Approaches to Study Cystic Fibrosis Pathophysiology Mark T. Clunes and Richard C. Boucher Abstract Mutation of the CFTR chloride channel was identified as the genetic basis of cystic fibrosis over 20 years ago; however, correlation of the pathophysiological changes occurring in CF lung disease with the mutation of a chloride channel is ongoing. The failure of innate lung defense in CF, and the subsequent cyclical microbial colonization of airways, explains the gross anatomical changes that occur in CF pathophysiology. However, ongoing research is focused on how the lack of the CFTR channel explains the failure of innate lung defense. Hydration status of the mucus blanket is key to understanding this link, and this series of chapters details the recent progress that has been made in understanding the interplay between ion transport activity and innate lung defense, and the initiation of CF lung pathophysiology. Key words: Airway surface liquid, defective ion transport, dehydration, mucus layer, respiratory infection, water transport.

1. Introduction Identification of the link between the failure of innate airway defense and defective ion transport has been the goal of cystic fibrosis research since CFTR was identified as a chloride channel (1). Although this area of research has been a focal point for 20 years, identifying causal events linking the failure of ion transport to development of respiratory infection/inflammation has been more difficult than anticipated. However, from data emerging over the past decade, the simplest scenario to link abnormal CF ion transport to CF lung disease is that CF epithelia have (1) reduced chloride and water transport capability and (2) poorly M.D. Amaral, K. Kunzelmann (eds.), Cystic Fibrosis, Methods in Molecular Biology 742, DOI 10.1007/978-1-61779-120-8_1, © Springer Science+Business Media, LLC 2011

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regulated sodium absorption, both leading to dehydrated airway surfaces. Dehydration of the mucus layer results in very different rheological properties from normal, with mucus becoming thickened and adhesive and resistant to the flow that is usually sustained by ciliary action in the airways. Static, adherent mucus then harbors pathogens within the lumen of CF airways. Indeed, adherent mucus ultimately stimulates the development of biofilms that are difficult to treat with inhaled antibiotics due to the obstruction to airflow. In addition, thickened mucus, which is a dense polymer gel, forms an effective barrier to neutrophil infiltration, allowing bacteria to thrive in the glycoprotein-rich environment (2). Infection then becomes persistent, and repeated cycles of inflammation culminate in airway remodeling and the development of severe obstructive and restrictive lung disease. Figure 1.1 outlines the mucociliary clearance mechanisms in place in normal lung and the changes that occur in CF lung.

Fig. 1.1. Airway surface dehydration and the development of mucociliary stasis. The left panel shows the normal situation where the airway surface liquid is maintained by a balance between chloride secretion, through CFTR, and sodium absorption, through the epithelial sodium channel (ENaC). By changing the mass of solute on the apical surface, the osmotic gradient for water secretion or absorption can be controlled and the airway surface and mucus kept hydrated. Hydrated mucus flows to the proximal airways, propelled by ciliary action. Right panel shows that when the chloride secretory pathway is absent, as in CF, the mass of NaCl in the apical domain is reduced. This is because there is no tonically active chloride secretory pathway and because the absorptive pathway is potentiated in the absence of CFTR. The consequence of the lack of CFTR is that no osmotic gradient is generated for water secretion. The airway surface becomes dehydrated and the mucus inspissated and adhesive, resulting in ciliary collapse and mucociliary stasis. Within the static mucus, bacterial biofilms form; in addition the dense polymer network (mucus), with a reduced mesh size, hinders neutrophil infiltration. Infection thus becomes established and proves near impossible to eradicate.

Approaches to Study Cystic Fibrosis Pathophysiology

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Effective disease management to date has focused on the amelioration of the symptoms of CF. Traditional therapies have focused on better nutrition and mechanical mechanisms that promote clearance of airway secretions, e.g., chest physiotherapy. Other therapies have centered on changing the rheological properties of mucus and controlling infection. The obstructive mucus plugs are the target of enzymes (e.g., DNase/RNase), reducing the viscoelastic properties of the DNA and promoting mucus clearance from the airways, either by ciliary action or by cough. In addition, aggressive antibiotic therapy, both parenteral and inhaled, has aimed to control existing infection and retard the development of further infection in “clear” areas of lung (3). More recently, therapies are being developed to treat the disease at more proximal steps in disease pathogenesis. For example, hypertonic saline delivered by nebulizer aims to rehydrate the mucus gel and restore clearance mechanisms (4, 5). Other therapies, e.g., denufosol, aim to re-direct salt transport from absorption to secretion, also hydrating airway surfaces. Further, as understanding of the molecular processes of CF pathophysiology have developed, whole new areas have opened up for targeted therapy. Therapies now aim to correct the basic defect in CF, namely defective CFTR processing and activation. 1.1. Localization of CFTR

The location of CFTR in the respiratory tree has been at the center of debate in CF physiology since initial studies, using polyclonal rabbit antibodies to CFTR, suggested that expression was dominant in the submucosal glands (6). This finding led to the conclusion that CF pathophysiology could be explained by submucosal gland dysfunction only. However, extrapolation of pathophysiological progression based on protein localization studies of epithelial channels traditionally has been difficult, as it is known that some channels, e.g., ENaC, are not abundantly expressed, nor detected immunohistochemically, despite robust physiological activity. So the question of how many channels are required to mediate significant physiological function remains unanswered. In addition, more recent studies, using sensitive mouse monoclonal antibodies with high-resolution laser confocal microscopy, have suggested that while submucosal glands can indeed express CFTR in the acinus, the ciliated duct is the predominant site of expression in the submucosal gland. Furthermore, the ciliated surface epithelial cells also routinely express CFTR both in the large and small airways (7). It seems, therefore, that CFTR is expressed and functional in both the proximal airway surface epithelium and the submucosal glands and that secretions from both sources may be defective in CF. In the small airways, where disease is initiated in CF, the ciliated cell selectively expresses CFTR.

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In Chapter 2, Silvia Kreda and Martina Gentzsch describe the latest methodologies used to immunohistologically visualize CFTR in human tissues and cultures, and in Chapter 6, Wine et al detail how intact whole submucosal glands can be prepared so that fluid secretion rates can be determined and their secretory regulation examined. Related to the issue of where CFTR is expressed is how CFTR is trafficked to the plasma membrane. Milder forms of CF pathophysiology are observed in which mutated CFTR channels are inserted into the membrane and allow some secretory function, e.g., R117H mutation (8). Yet, with F508, very little, if any, protein actually trafficks to the plasma membrane under normal circumstances. Under experimental low-temperature conditions, F508 CFTR can be trafficked to the membrane (9), and is known to have functional activity once in the membrane, albeit lesser than wild type (10). In addition, even when F508 CFTR is experimentally over-expressed in heterologous systems, its turnover in the plasma membrane is rapid, suggesting that some form of mechanism exists that recognizes mutated protein in the plasma membrane and responds by removing that protein (11). Understanding of how CFTR is trafficked to the membrane and the pathways that control and regulate this trafficking will undoubtedly provide a rich source of targets for pharmacological intervention. To this end, Martina Gentzsch and Jack Riordan have produced epitope-tagged forms of CFTR that can be easily traced throughout the lifetime of CFTR’s existence in the cell, from its formation in ER to its insertion into the membrane and subsequent turnover. This invaluable tool allows the natural history of F508 CFTR to be observed, and the importance of this tool is highlighted by research to discover “corrector” drugs that aim to deliver F508 protein to the membrane and potentiator drugs to stabilize and activate the protein once in the membrane. 1.2. Ion Transport and Airway Surface Liquid

The developmental aspects of ion transport in the CF fetal/neonatal lung are worth noting. CF is not widely regarded as a cause of intrauterine death. This is surprising, since lung development necessitates the secretion of fluid into the lung to generate the transmural pressure to stimulate lung growth (12). This fetal lung secretory process is apparently undisturbed in CF. For example, although problems that lead to a paucity of fluid in the fetal lung are recognized to produce small, underdeveloped lungs, e.g., oligohydramnios (13), this pathology is not generally a feature of CF. Although fetal CF genotyping is rare, only subtle changes in respiratory morphology have been reported in CF fetuses (14). In itself, this observation suggests that fetal CF lungs have the ability to secrete fluid, and that this secondary mechanism is not retained in the adult CF lung, or that mutated CFTR is handled very differently by the cell during embryonic development. An “alternative” chloride secretory pathway has


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been well defined as the P2Y2 receptor-mediated, calciumactivated chloride pathway [CaCC (15)] and is known to be active in the fetal (16) as well as the adult lung. Consequently, it is possible that CaCC, or as yet unrecognized Cl− channels, mediate fetal lung liquid secretion. Of note, the CaCC secretory pathway has been focused upon as a therapeutic target for the treatment of CF. These ionic conduction pathways are important, since transepithelial water flux is governed by the net rate of Na+ absorption vs. the net rate of Cl− secretion. It is the transepithelial secretion of chloride ions into the airway lumen that provides the electrical driving force (transepithelial potential difference) for Na+ to follow. Thus, NaCl is secreted onto the airway surface and provides an osmotic driving force for water transport. Two major routes for chloride secretion are utilized by the postnatal airway (1): the cystic fibrosis transmembrane conductance regulator (CFTR) and (2) the calcium-activated chloride conductance (CaCC). Although the dominant baseline secretory activity present in airway epithelial cells is the PKA-regulated CFTR Cl− channel, the loss of which results in dehydrated airways and CF lung disease, the CaCC pathway remains intact in CF and represents a second route for chloride secretion that can also be stimulated via a number of mechanisms. Salt and water absorption on the other hand is achieved solely by the activity of the apically located Na+ channel ENaC. The electrogenic transepithelial transport of Na+ from the apical surface liquid into the basolateral space provides the electrical driving force for paracellular anion transport; thus, NaCl is transported from the airway surface liquid to the basolateral interstitial space. The generated osmotic gradient produces transepithelial water absorption. Thus, fluid transport, and hence airway surface volume, is determined by the ratio of basolateral-to-apical Cl− secretion to apical-to-basolateral Na+ absorption. 1.3. Ion Transport In Vivo/In Vitro

The bioelectric properties of airway epithelia have been hard to assess in vivo because of the complex three-dimensional architecture of the respiratory tree. Perhaps one of the easiest in vivo measurements, and one that has now become diagnostic for CF, is the measurement of the airway transepithelial electric potential difference (PD) (17). The CF nasal epithelium exhibits a very high basal potential difference that is largely amiloride sensitive, indicating that the basal bioelectric properties are dominated by sodium hyperabsorption (18). Nasal PD measurements are a wonderful tool to study ion channel function in vivo. However, the nasal epithelium is not necessarily fully representative of the transport processes occurring deeper in the respiratory tree, particularly at the bronchiolar level where CF disease begins. In vivo PD measurements in the small airways would be exceedingly difficult, but the development of human-cultured bronchial epithelial cells

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has allowed the assessment of bioelectric and fluid handling properties of airway cells from all regions of the respiratory tree in a culture system. Human bronchial epithelial cells have been successfully cultured and their bioelectric properties studied in Ussing chambers for many years. However, it is the constant requirement for human tissue that limits this approach. Refinement of the culture methods so that cells more closely resemble native human tissue and extension of the useful life of cultured cells has been a priority in this field. Sophie Crespin et al (Chapter 10) examine the markers for well-differentiated human airway epithelial cells and discusses how cells can be maintained and studied in long-term culture (19). 1.4. Fluid Transport

The development of bronchial epithelial cell cultures has also been instrumental in gaining insight into the regulation of airway surface liquid volume. Use of large molecular weight fluorescent dyes (e.g., dextran-conjugated Texas Red) that are cell impermeable allows the airway surface liquid depth to be monitored using Z stack confocal microscopy. The advantage of this methodology is that liquid height can be resolved down to the micrometer range and, thus, ASL height can be monitored over time in live cultures. The permeable supports on which cultures are grown allow the epithelium to polarize and independent pharmacological manipulation of basolateral or apical domains is possible. This methodology has been used extensively to assess the function of both CFTR-mediated fluid secretion and CaCC-mediated fluid secretion in CF cultures. Robert Tarran and Erin Worthington (Chapter 5) discuss the latest developments in the assessment of airway surface liquid volumes using this technique.

1.5. Fluid and Mucus

Traditionally it had been thought that the mucus floated on the periciliary layer of fluid termed sol. In consequence, it was assumed that as the fluid depth diminished, the mucus would compress the cilia and retard their action, and indeed this is true [for review, see (20)]. It was also assumed that too much fluid would allow the mucin to lift away from the ciliary tips and that mucociliary transport would suffer due to a loss of traction by the cilia on the mucus blanket. Indeed, this may not be true. Mucus is a polymer gel; some mucins are tethered to the epithelial surface, while others are truly secreted into the luminal space; the hydration state of the mucus is critical for innate defense. However, it is now known that the polymer gel can swell and accommodate “extra” airway surface liquid, giving the gel a looser formation and actually increasing mucociliary transport rates (21). It is a paucity of fluid and the development of viscous, adherent mucus that precipitate CF pathophysiology. Günthar Hansson and Kristina Thomsson (Chapter 8) examine the latest proteomic


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methods used to identify the various mucins present in mucus and how these mucins can be adequately quantified (22). This is an essential step for CF pathobiology so that we can understand the exact solvent requirements for effective mucociliary clearance. 1.6. Mechanical Sensitivity – The CaCC Pathway

The lung is a dynamic organ, and both airways and alveoli are stretched during the respiratory cycle. The mechanical stimulation of airway cells, either by physical deformation or by flowinduced shear stress across the cells, results in the release of ATP [review (23)] and possibly other ATP/UTP metabolites into the lumen. ATP concentrations rise in the airway surface liquid with mechanical stimulation, along with the concentration of ATP metabolites such as ADP, AMP, and adenosine, generated by ecto-enzymes on the airway surface (24). ATP, an agonist of apical P2Y2 receptors, increases free intracellular calcium ([Ca2+ ]i ), leading to activation of CaCC and net transport of NaCl into the airway surface liquid, providing the osmotic force for water transport. In part, this mechanism explains why mechanical stimulation of the lungs results in “bursts” of secretion. Indeed, many mechanical “stimulants” are used in CF therapy, including airway clearance techniques, and percussion, either manually or with a mechanical “vest,” to assist airway mucus clearance. [Ca2+ ]i , in addition to activating CaCC, is an important modulator of mucin secretion and ciliary beat frequency and exhibits adaptation to inflammatory responses. [Ca2+ ]i , then, is important in the regulation of innate defense, and in the early 1990s, CF researchers took advantage of the newly developed tools for monitoring [Ca2+ ]i that had been developed by Roger Tsien (25). In Chapter 7, Carla Ribeiro examines these techniques and focuses on how the unique problems that airway researchers faced in [Ca2+ ]i measurements were overcome. [Ca2+ ]i was initially measured in dissociated cells that were grown or attached to a glass substrate. In this way, cells were easier to load with fluorescent dye (because they were rounded up), and they were positioned on a substrate with the right optical qualities to allow shortwavelength excitation light to reach the dye. This approach was suboptimal for airway research, since airway cells exhibit a polarized nature in which the basolateral and apical spaces are distinct. Long working distance fluorescent lenses, coupled with the use of dyes with a high quantum yield, like fura-2, allowed studies of polarized epithelia. Thus, airway cells could be grown on permeable supports as a sheet of epithelial cells and the basolateral and apical spaces could be independently accessed experimentally. This advance was important, since pharmacological interventions were shown to be able to selectively target the apical domain vs. the basolateral domain in isolation, and we now know that the resident receptor populations differ significantly between these domains (26) as well as calcium handling (27).

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1.7. Pathway Interaction – CaCC, CFTR, and ENaC

An increase in intracellular cAMP levels leads to an increase in CFTR activity and produces chloride and fluid secretion. To maximize this secretion, it is now clear that CFTR directly regulates the activity of ENaC and switches off absorption during activation of its secretory activity. In CF epithelia, this lack of antagonism of ENaC by CFTR is responsible for the hyperabsorption observed in CF airways. Early work with the patch clamp technique demonstrated this antagonistic nature between CFTR function and ENaC function (28) and suggested that proximity of the channels to one another was significant, since the area included in a single patch is less than 1 μm in diameter. This study suggested association between the channels, and later studies using FRET (Fluorescent Resonant Energy Transfer) have confirmed this close association between CFTR and ENaC. FRET studies require the expression of tagged CFTR and ENaC. For example, CFTR is tagged with ECFP (enhanced cyan fluorescent protein), while ENaC is tagged with EYFP (enhanced yellow fluorescent protein). Short-wavelength excitation of ECFP results in fluorescent emission of a longer wavelength that is suitable for excitation of EYFP and the resulting longer wavelength fluorescence of EYFP can then be measured. Very close association between the fluorescent molecules is required ( 300 cm2 .

2.2.2. Culture of Cell Lines Heterologously Expressing CFTR

BHK-21 cells stably expressing CFTR, F508del, and GFP- or extracellularly HA epitope-tagged variants [Extope-CFTR; (6)] are grown in DMEM/F12, 5% FBS, and 500 μM methotrexate at low confluence on glass bottom microwell dishes (MatTek Corporation, USA) or BD BioCoat collagen-coated culture slides (BD Biosciences, USA) (see Note 1). In cell lines stably expressing CFTR, the levels of CFTR protein expression can be increased

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by addition of 2 mM sodium butyrate to the growth medium for 8–16 h before the experiment. 2.3. Frozen and Paraffin-Embedded Primary Cultures

1. Well-differentiated primary cultures are embedded in OCT and cryopreserved at –80◦ C. Cryosections are performed as above. 2. Well-differentiated primary cultures are fixed in buffered formalin (Fisher, USA) overnight at room temperature and then embedded in paraffin (Fisher, USA). Thin sections (∼5 μm thick) are generated on plus glass slides.

2.4. Mouse Tissue Specimens

2.5. Immunostaining Reagents

Excised mouse lungs and tracheas are quickly embedded in OCT and cryopreserved at –80◦ C. Cryosections are produced as described above. 1. Fixation. 4% paraformaldehyde solution freshly prepared by diluting 16% paraformaldehyde purchased from Electron Microscopy Sciences (Hatfield, PA; stored protected from light, airtight, at 4◦ C) in phosphate buffer solution with 100 μM each of calcium and magnesium (PBS). Methanol and acetone were obtained from Fisher (USA). 2. Permeabilization. 100% methanol (HPLC quality; Fisher, USA) or alternatively 0.1% Triton X-100 prepared fresh by diluting 10% Triton X-100 (Pierce, Rockford, IL; stored airtight, at 4◦ C) in PBS or 0.1% saponin prepared fresh by diluting 10% saponin (Sigma, USA) in PBS. 3. Blocking. 10–20% non-immune normal serum (NS) of secondary antibody species (Jackson ImmunoResearch Labs, West Grove, PA) or alternatively 1% fatty acid-free bovine serum albumin (BSA) of RIA quality (Sigma, USA). Serum dilution is freshly prepared. BSA solution is prepared from a 5% BSA/PBS stock solution aliquoted and stored at –20◦ C. 4. CFTR monoclonal antibodies. CFTR antibodies (clones 528, 769, 596, and 570) were developed and provided by Dr. John R. Riordan (University of North Carolina at Chapel Hill) (2, 3, 5, 9). 5. Other antibodies and probes. Ezrin mouse monoclonal antibody is from BD Biosciences (San Jose, CA), tubulin rat monoclonal to label cilia is from Chemicon (Temecula, CA), rabbit polyclonal antibody against MUC5AC to identify goblet cells/mucin granules is a kind gift from John Sheehan (UNC), and fluorescently labeled phalloidin to label actin cytoskeleton is from Molecular Probes (Eugene, OR). Anti-HA antibodies HA.11 (clone 16B12; Covance, USA) and 12CA5 (Roche, Germany) are used to visualize Extope-CFTR.

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6. Non-immune mouse IgG and fluorescently labeled secondary antibodies are from Jackson ImmunoResearch Labs and Molecular Probes (Invitrogen, USA), respectively. 7. Vectashield mounting medium containing DAPI, to label nuclei, is purchased from Vector Labs (Burlingame, CA) or alternatively SlowFade Gold antifade reagent is purchased from Molecular Probes (Invitrogen, USA). 8. Epitope retrieval solution is from DAKO USA (Carpinteria, CA) and used following manufacturer’s instructions to unmask CFTR epitopes in paraffin-embedded specimens. 9. Vector M.O.M. Basic Kit (Vector Labs) for immunostaining of mouse tissues with antibodies produced in mouse and Texas Red-labeled streptavidin (Jackson ImmunoResearch Labs) to detect mouse monoclonal antibody immunostaining. 10. Wet chamber. A plastic box with a fitted lid containing several plastic pipettes organized in parallel to hold glass slides and a clean piece of wet paper on the bottom to provide humidity. 11. Histology pen is purchased from Fisher (USA). 12. Glass cover slips number 1.5 are manufactured by Corning USA (Fisher, USA). 13. Nail polish to seal mounted specimens. 14. Leica laser confocal microscopy system SP2 AOBS with an upright microscope DM-RXA2 containing a 16× PL Fluotar (NA 0.5), 40× Apochromat (NA 1.25–0.75, oil), 63× PlanApo (NA 1.4–0.6) Leica lenses, and DIC/Nomarski illumination. The scanning system is equipped with a highprecision galvanometer stage for real-time xz scanning and four independent lasers, namely UV 351/364 nm for DAPI, Argon 488 nm for FITC and Alexa Fluor 488, solid state diode pump 561 nm for Texas Red and Alexa Fluor 568, and HeNe 633 nm for CY5 and Alexa Fluor 633. Zeiss LSM 510 confocal laser scanning microscope with high-precision galvanometer stage, a Plan Apochromat 63× lens (NA 1.4, oil), DIC/Nomarski illumination, and lasers Argon 488 nm, HeNe 543 nm for Texas Red and Alexa Fluor 568, and HeNe 633 nm.

3. Methods In this section we describe immunolocalization of endogenous and heterologously expressed CFTR protein in tissues and cell cultures. These techniques can be adapted to any type of cell culture and tissue. Protocols for fresh, frozen, and

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formalin-fixed/paraffin-embedded specimens are illustrated. We do not recommend the use of paraffin-embedded specimens because of variable and deficient preservation of CFTR protein levels and/or CFTR reactivity to antibodies. Instead, we use fresh or quickly frozen specimens preserved by controlled, mild fixation, since fixation can affect the capacity of antibodies to recognize CFTR (4). We use the CFTR monoclonal antibodies developed by Dr. John R. Riordan (UNC-CH) (2, 3, 5, 9). CFTR antibodies from other sources produce variable results in immunolocalization studies (2–4, 10, 11). We illustrate co-staining of CFTR antibodies with antibodies or probes reacting against cellular markers for identification of cell types or intracellular compartments to facilitate the interpretation of CFTR localization studies. Our protocols use fluorescence as the detection method and laser confocal microscopy for data collection and analysis. Confocal microscopy has optical advantages over epifluorescence microscopy, the former being more reliable in discriminating different fluorescent signals in co-staining experiments, identifying subcellular localization of fluorescent signals, and comparing localization of a fluorescent probe among different specimens (12). We will also discuss techniques for immunostaining of heterologously expressed human CFTR in cell cultures and mouse tissues. The high levels of CFTR protein expression in human heterologous cells allow immunostaining to be easier than in native cells. However, in mouse tissues, a different approach needs to be used to overcome the use of mouse monoclonal CFTR antibodies on mouse tissues. Because GFP-tagged CFTR can be used for trafficking studies, it has been included in this chapter. However, we favor the use of Extope-CFTR, which refers to CFTR with an externally accessible HA epitope inserted into an extended, modified extracellular loop 2 that contains sequences from the first and fourth extracellular loops to expose the HA epitope (6, 7). Extope-CFTR matures and functions like unmodified wild-type CFTR (6) and permits selective visualization of surface CFTR and chasing of labeled CFTR during internalization for monitoring of CFTR protein recycling in intact cells (6, 7). 3.1. Immunofluorescence Staining of Endogenous CFTR in Cryosections of Human Tissues

Cryosections of frozen freshly excised airway, rectal, or skin tissues mounted on glass slides can be processed for CFTR immunolocalization using the following protocol: 1. Fixation. Frozen tissue slides are air-thawed and promptly fixed with 4% paraformaldehyde/PBS solution for 5 min at room temperature (see Note 2). Fixative is eliminated and slides are washed twice for 5 min at room temperature. 2. Permeabilization. Tissue slides are rinsed with PBS and incubated with 100% ethanol for 2 min at –20◦ C (see Note 3). 3. Blocking. Slides are air-dried and tissue specimens encircled using a histology pen (see Note 4). Tissue wells are rinsed

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once with PBS and incubated in 10% normal serum/PBS blocking solution for at least 30 min at room temperature (see Note 5) using a wet chamber (see Note 6). 4. First antibody incubation. Incubation with CFTR antibodies, non-immune mouse IgG, and other antibodies is performed overnight at 4◦ C (see Note 7). Antibodies and IgG are diluted in blocking solution; CFTR antibodies and IgG are used at matching concentrations of 5–10 μg/ml, and ezrin, tubulin, and MUC5AC antibodies are employed at 0.5–1 μg/ml. In co-staining experiments, CFTR and other marker antibodies are added together at the specified concentrations. 5. Washes. Antibody solutions are discarded and tissues washed twice with PBS for 5 min at room temperature. 6. Secondary antibody incubation. Fluorescently labeled secondary antibodies are diluted in blocking solution at the time of the experiment, at the concentration recommended by the manufacturer, and added for 1 h at room temperature, protected from light. In co-staining experiments, all secondary antibodies are added together (see Notes 8 and 9). Fluorescently labeled phalloidin can be added at this step at 0.5–1 μM concentration. 7. Washes. Antibody solutions are discarded and cultures washed three times with PBS for 5 min at room temperature, protected from light. 8. Mounting. Most of the liquid should be eliminated without air-drying the tissue sections (see Note 6). A glass cover slip with a diameter larger than that of the tissue section is placed on a piece of absorbent paper and a drop of mounting medium is added on it. The glass slide is inverted with the tissue facing down and is carefully laid over the cover slip, starting from one side to avoid the formation of air bubbles. The excess mounting medium is allowed to drain into the paper for a few seconds without pressing on the glass (see Note 10). Any liquid on the glass is eliminated with a vacuum-attached Pasteur pipette and the cover slip is sealed with nail polish. It is stored at 4◦ C, protected from light. 9. Confocal microscopy and image analysis. Confocal microscopy analysis is performed by xy scanning using the appropriate lasers (see Note 11), a 63× lens, and Nomarski illumination (see Note 12). Parameters should be maintained constant throughout the analysis. Imaging analysis and montage are performed using Leica and Adobe PS software, respectively (see Note 13). Examples of the results obtained using this protocol are illustrated in Figs. 2.1a, b and 2.2a.

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3.2. Immunofluorescence Staining of Endogenous CFTR in Formalin-Fixed, Paraffin-Embedded Human Tissues

1. Formalin-fixed, paraffin-embedded tissues mounted on glass slides (see Note 14) are deparaffinized by sequentially immersing glass slides in xylol and 100, 95, and 70% alcohol solutions.

3.3. Immunofluorescence Staining of Endogenous Human CFTR in Fresh Well-Differentiated Primary Airway Epithelial Cultures

1. Fixation. Well-differentiated bronchial epithelial cultures are rinsed twice with PBS and fixed by adding 4% paraformaldehyde/PBS solution to both the apical and basolateral sides. Fixation is for 5 min at room temperature (see Note 2). Fixative is eliminated and washed twice for 5 min at room temperature.

2. Samples are air-dried and subjected to epitope retrieval with DAKO (pH 6) solution using a microwave for 1 min at full power followed by 10 min at 10% power. Samples are cooled to room temperature, rinsed with PBS, and subjected to the blocking and subsequent steps as described in Section 3.1 for frozen tissues.

2. Permeabilization. Cultures are incubated with 0.1% Triton X-100/PBS solution on both sides for 20 min at room temperature (see Note 3). 3. Blocking. Cultures are rinsed twice with PBS and incubated apically with 100 μl of 1% BSA/PBS blocking solution for at least 30 min at room temperature (see Notes 5 and 6). 4. First antibody incubation. Incubation with CFTR antibodies, non-immune mouse IgG, and other antibodies is performed as described in Section 3.1 for frozen tissues. A 12-mm insert requires 100 μl of antibody solution to cover the apical side. The basolateral side may be bathed in 100–200 μl of PBS (see Note 7). 5. Washes and secondary antibody incubation. Washes after the first and secondary antibodies, and incubation with fluorescently labeled secondary antibodies and fluorescently labeled phalloidin are performed as described in Section 3.1 for frozen tissues (see Notes 8 and 9). 6. Mounting. The membrane supporting the culture is carefully separated from the plastic cup insert using a surgical knife and placed flat on a glass slide with the help of forceps. A minimal amount of liquid should be present in the insert before starting this operation. With gentle outward movements, the edges of the cultured membrane are combed with a surgical knife to dislodge cell sheets; the center of the culture should remain intact. A glass cover slip with a diameter larger than that of the cultured membrane is placed on a piece of absorbent paper and a drop of mounting medium is added on it. The glass slide is inverted with the cell culture facing down and is carefully laid over the cover slip,

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starting from one side to avoid the formation of air bubbles. The excess mounting medium is allowed to drain into the paper for a few seconds without pressing on the glass (see Note 10). Any liquid on the glass is eliminated with a vacuum-attached Pasteur pipette and the cover slip is sealed with nail polish. It is stored at 4◦ C, protected from light. 7. Confocal microscopy and image analysis. For specimen imaging and analysis, we use a Leica laser confocal microscopy system SP2 AOBS with an upright microscope DM-RXA2. Specimens are imaged using the appropriate lasers (see Note 11) and a 63× lens by xz (cross section) scanning in the center of the cultures, where the epithelial architecture is left intact. Cells scraped off the membrane support are xy (in face) scanned using simultaneous DIC and fluorescence channels (see Note 12). The conditions for scanning are kept constant throughout the studies. Imaging analysis and montage are performed using Leica and Adobe PS software, respectively (see Note 13). An example of the results obtained using this protocol is illustrated in Fig. 2.1c. 3.4. Immunofluorescence Staining of Endogenous Human CFTR in Frozen or Formalin-Fixed/ Paraffin-Embedded, Well-Differentiated Primary Airway Epithelial Cultures

1. Frozen cell culture sections mounted on glass slides (see Note 15) are air-thawed and promptly fixed and processed for immunostaining following the protocol described in Section 3.1 for frozen tissues. 2. Formalin-fixed, paraffin-embedded culture sections mounted on glass slides are deparaffinized and processed for immunostaining as described in Section 3.2 for paraffinembedded tissues (see Note 14).

3.5. Immunostaining of CFTR Heterologously Expressed in Cell Cultures Using Anti-CFTR Antibodies Detecting Intracellular Epitopes

Cell cultures heterologously expressing human CFTR are rinsed twice with PBS and processed for immunostaining following the protocol described in Section 3.1 for frozen human tissues (see Notes 16, 17, and 18). An example of results obtained using this protocol is shown in Fig. 2.3a, c.

3.6. Visualization of CFTR Pools at the Plasma Membrane and in Endocytic Compartments, Using an Extracellular Epitope Tag

Surface CFTR is labeled with HA antibodies at 4◦ C to avoid internalization. 1. Wash cells (e.g., BHK-21 cells) stably expressing ExtopeCFTR three times with PBS. 2. Pre-cool cells for 15 min on ice in a cold room at 4◦ C. 3. Label cell surface Extope-CFTR for 30 min on ice in a cold room with anti-HA 16B12 raw ascites diluted 1:500 in PBS with 1% BSA (see Note 19).

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4. Wash cells four times with ice-cold PBS. If you intend to visualize surface CFTR, proceed to Step 5. For visualization of endocytosed CFTR, proceed to Step 8. 5. Fix cells with cold 4% paraformaldehyde for 10 min (see Note 20). 6. Block and incubate with goat anti-mouse IgG Alexa Fluor 488 antibody. 7. Perform washes, mounting, and confocal microscopy analysis in a Zeiss LSM 510 system as described in Section 3.1 for frozen tissues. 8. For visualization of endocytosed CFTR, add warm growth media to cells after Step 4 and reincubate cells at 37◦ C in a cell incubator for the indicated times. 9. Rinse cells twice quickly with PBS. 10. Fix cells in 4% paraformaldehyde in PBS at room temperature for 10 min (see Note 16). 11. Wash three times for 10 min with PBS at room temperature. 12. Permeabilize cells with 0.1% Triton X-100 in PBS for 10 min at room temperature (see Note 17). 13. Wash three times for 10 min. 14. Block at least for 1 h with blocking buffer. 15. Perform incubation with fluorescently labeled secondary antibodies, washes, mounting, and confocal microscopy analysis as described in Step 7. An example of results obtained using this protocol is shown in Fig. 2.3e. 3.7. Visualization of CFTR Recycling from an Intracellular Compartment to the Plasma Membrane, Using an Extracellular Epitope Tag

To study recycling of Extope-CFTR, HA antibodies bound to surface CFTR are removed by a low pH wash resulting in label remaining exclusively on CFTR in endocytic vesicles. CFTR that recycles to the surface after reincubation is then visualized on fixed, but not permeabilized cells. 1. Label cells (e.g., BHK-21) expressing Extope-CFTR by addition of monoclonal mouse anti-HA antibody 12CA5 to the growth medium for 10 min at 37◦ C. 2. Quickly remove 12CA5 antibody bound to cell surface pools of Extope-CFTR by rinsing cells twice for 30 s with ice-cold PBS (pH 3.7) (PBS + HCl), followed by a wash with PBS (pH 7.4). Fix one sample at this step with 4% paraformaldehyde to verify complete removal of apical label. 3. Reincubate cells in pre-warmed media for 5 min at 37◦ C. Fix cells with 4% paraformaldehyde, block, and incubate with goat anti-mouse IgG Alexa Fluor 488 antibody as described in Section 3.6 to visualize CFTR that has been recycled to the cell surface (see Note 20).

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4. Perform washes, mounting, and confocal microscopy analysis as described in Section 3.6. 3.8. Imaging of Green Fluorescent Protein-Tagged CFTR

Green fluorescent protein-tagged CFTR can be visualized in live cells or in previously fixed cells. Live cells are best observed in dye-free, HEPES-buffered growth medium (see Note 21) (see Fig. 2.3b).

3.9. Immunofluorescence Staining of Transgenic Human CFTR in Frozen Mouse Tissues

1. Frozen tissue sections are fixed in cold 100% acetone (see Note 22). 2. Immunostaining with antibodies against human CFTR is performed using the Vector M.O.M. Basic Kit (as per manufacturer’s instructions) for optimized immunodetection in mouse tissues, using mouse-raised antibodies. Briefly, CFTR antibodies are incubated at 10 μg/ml (prepared in M.O.M. diluent) for 60 min at room temperature. M.O.M. biotinylated anti-mouse IgG reagent is incubated for 10 min at room temperature. Texas Red streptavidin is incubated at 15 μg/ml (prepared in PBS) for 10 min at room temperature. 3. Mounting and confocal microscopy analysis are performed by xy scanning as described in Section 3.1 for human tissues.

4. Notes 1. Different cell lines have different requirements for none or specific coating for the glass substrata and growth media. 2. Fixation is critical for effective immunoreactivity with CFTR monoclonal antibodies. Paraformaldehyde dilution is freshly prepared from a high-quality source. Long fixation times may require an epitope retrieval step before incubation with CFTR antibodies and is not recommended. 3. Alternatively, permeabilization can be performed with 100% methanol or ethanol for 5 min at –20◦ C; when using this permeabilization, fixation can be done with 2% paraformaldehyde. Alcohol permeabilization usually works better in preserving cellular integrity in freshly excised frozen tissues. 4. The use of a hydrophobic histology pen to draw wells encircling the tissue specimens allows the use of reduced amounts of immunostaining reagents. If more than one tissue section is mounted on a glass slide, each section can be incubated with a different antibody if separated in different pen-drawn wells. Tissues should be sufficiently separated to

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ensure no cross-contamination between wells using different antibodies. 5. RIA-quality BSA is recommended because it is mostly devoid of other serum components. Immunostaining of cultures can be achieved using a low-concentration BSA solution for blocking non-specific antibody binding. In contrast, tissues may require better blocking. Thus, a solution prepared with 10–20% of non-immune (normal) serum from the secondary antibody species, with/without the addition of 3% BSA, usually works better than BSA alone in airway tissues. 6. Cultures should not be allowed to dry during the immunostaining proceeding to avoid staining artifacts. For specimens on glass slides, use a wet chamber, and for cell cultures grown on plates, replace the lid after each incubation step. 7. Incubation with CFTR antibodies overnight at 4◦ C yields a better signal/background ratio than does incubation for 2 h at room temperature. This is particularly important in airway tissues, due to the relative low expression of the CFTR protein vs. the high capacity of non-specific antibody binding of the different tissue structures. 8. Secondary antibodies should have minimal cross-reactivity with human proteins and the secondary antibody species. Fluorescently labeled Fab or F(ab )2 antibody fragments are very useful in avoiding cross-reactivity, particularly in airway tissues undergoing inflammation and mucous metaplasia. 9. The part of a compound that emits fluorescence is a fluorophore. In co-staining experiments, the selection of fluorophores labeling antibodies and phalloidin must be based on the excitation/emission characteristics of the molecules such that they do not interfere with each other in the microscope analysis. Fluorophores with broad excitation/emission characteristics should be avoided when costaining CFTR with other cellular markers. Information about fluorescent molecules can be accessed at Molecular Probes and Jackson ImmunoResearch Labs. 10. Mounting medium thickness is critical when using low working distance microscopy immersion lenses, because too much medium will render poor focusing. In contrast, too little medium will favor the generation of air bubbles and will compress the cultures between both glasses, altering the epithelial architecture. 11. Confocal microscopy is desired for the analysis of CFTR localization. The thin optical sectioning produced by laser

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confocal scanning ensures accurate spatial collection of immunostaining signal necessary to localize CFTR protein relative to other cellular structures or markers (e.g., ciliated cells, plasma membrane, and intracellular vesicles). Moreover, many laser confocal scanning systems are equipped to adjust to narrow excitation and emission bandwidths to optically discriminate different fluorophores present simultaneously in co-localization experiments. However, the success of co-localization experiments relies on selecting correctly a set of fluorophores that will be accurately detected and discriminated by the microscopy system. Laser confocal microscopy systems equipped with a galvanometer stage allow real-time xz scanning of epithelial cultures to image a cross section of the cells, as opposed to image reconstruction with specific software of a timeconsuming, high-resolution xy multi-focal stack. 12. Differential interference contrast (DIC) or Nomarski illumination produces images of detailed cell structure for morphology and localization analyses and is recommended for imaging immunolocalization data in tissues. 13. Unbiased image collection and analysis of CFTR immunostaining are crucial, mainly in specimens where CFTR localization may be differently affected (i.e., normal vs. CF airway epithelia). Thus, studies should be performed blind to the identity/phenotype of the specimens, and conditions used in image collection and analysis should be maintained constant throughout. Similarly, alterations introduced in the images for presentation or publication purposes should affect identically all the images. Image alteration should be avoided or kept to a minimum and should be done only for aesthetic or didactic reasons without affecting the integrity of the scientific data. 14. Formalin-fixed, paraffin-embedded, well-differentiated airway epithelial cultures or tissue specimens yield wellpreserved cell specimens that can be stored for long periods of time. However, most CFTR antibodies require strong epitope retrieval techniques in these specimens, with varying results; this means, the levels of CFTR immunoreactivity may misrepresent actual CFTR protein levels in the specimens, and comparison between different specimens may be difficult. Epitope retrieval techniques also often damage fine cellular architecture and preclude the use of high-power magnification/resolution microscopy techniques. Moreover, analysis of co-localization studies may be affected by blemished cell architecture integrity produced by the epitope retrieval technique. Therefore, caution should be exercised using formalin-fixed,

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paraffin-embedded specimens for CFTR immunostaining analysis. Culture and tissue specimens for long-term histology collections are usually preserved using the formalinfixation/paraffin-embedding techniques. However, cytological preservation, and therefore CFTR protein integrity, is affected by the timing between specimen collection and fixation, fixation method (e.g., fixative type and concentration, duration of fixation), paraffin-embedding method, and storing conditions (i.e., time, temperature, and humidity), which vary through time and between different laboratories. Thus, formalin-fixed, paraffin-embedded specimens are not suitable for studies aimed to characterize localization of CFTR or compare CFTR immunostaining levels between different specimens. These specimens should never be used as the primary source for a CFTR immunostaining study! 15. Frozen slides of well-differentiated airway epithelial cultures are very useful because many slices can be obtained from one culture. However, freezing and cutting epithelial cultures grown on membrane supports require an experienced hand. Often, cultures are damaged during the freezing or cutting steps and are not suitable for CFTR immunolocalization studies. 16. Alternatively, non-polarized cells may be fixed with ice-cold methanol at –20◦ C for 10 min. The permeabilization step is not necessary, since alcohol functions as both fixative and permeabilizing agent. 17. Alternatively, cells fixed with 4% paraformaldehyde may be permeabilized with 0.1% saponin in PBS for 1 h at 4◦ C. 18. For cells overexpressing CFTR, immunostaining may be performed using raw ascites fluid containing anti-CFTR monoclonal antibodies 570, 596, 528 diluted 1:500 and incubated for 1 h at room temperature or 1:1000 and incubated overnight at 4◦ C. 19. Anti-HA antibody HA.11 (clone 16B12; Covance, USA) is generally used to visualize the extracellular tag in ExtopeCFTR. HA.11 anti-HA antibody binds to epitope in Extope-CFTR even at pH 3–4 and is therefore not useful for studies in which the antibody has to be removed from the epitope. In contrast, anti-HA antibody 12CA5 (Roche, Germany) can be removed from cell surface Extope-CFTR by an acidic wash and is consequently used in recycling studies intended to monitor internalized Extope-CFTR. 20. Do not use MeOH for fixation, as this maneuver will permeabilize the cells. Similarly, do not permeabilize the cells

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with detergent before blocking, as this will lead to visualization of intracellular Extope-CFTR pools. If desired, intracellular pools of CFTR can be visualized in parallel permeabilized samples; however, keep these slides away from slides that will not be permeabilized to avoid contamination with detergent. 21. When observing live cells, avoid bleaching of the sample, alkalinization of the culture medium, and also excess heating by laser exposure. 22. Acetone fixation is recommended by the manufacturer of the M.O.M. kit to minimize background staining.

Acknowledgments The authors gratefully thank the CF and non-CF volunteers for their tissue specimen donation; Richard Boucher for his comments, John R. Riordan and John Sheehan for their generous gifts of CFTR antibodies and MUC5AC polyclonal antibody, respectively; James Yankaskas and Marcus Mall for human tissue specimen collection; Scott Randell and Leslie Fulcher for providing the primary airway epithelial cells, Kim Burns, Tracy Barlotta, and Donald Joyner for their expert technical assistance; Lisa Brown for editing this manuscript; Michael Chua and Neal Kramarcy for microscopy assistance; and the UNC M. Hooker Microscopy Facility for making accessible their microscopes. This work is supported by CFF grants GENTZS04G0 and GENTZS07G0 (MG) and KREDA01I0 (SMK), the Mary Lynn Richardson Fund (SMK), and NIH Grants HL34322 and HL 51818-06A1 (SMK). References 1. Kartner, N., Augustinas, O., Jensen, T. J., Naismith, A. L., and Riordan, J. R. (1992) Mislocalization of DF508 CFTR in cystic fibrosis sweat gland. Nat Genet 1, 321–327. 2. Mall, M., Kreda, S. M., Mengos, A., Jensen, T. J., Hirtz, S., Seydewitz, H. H., et al. (2004) The DeltaF508 mutation results in loss of CFTR function and mature protein in native human colon. Gastroenterology 126, 32–41. 3. Kreda, S. M., Mall, M., Mengos, A., Rochelle, L., Yankaskas, J., Riordan, J. R., et al. (2005) Characterization of wild-type

and {Delta}F508 cystic fibrosis transmembrane regulator in human respiratory epithelia. Mol Biol Cell 16, 2154–2167. 4. Claass, A., Sommer, M., de, J. H., Kälin, N., and Tümmler, B. (2000) Applicability of different antibodies for immunohistochemical localization of CFTR in sweat glands from healthy controls and from patients with cystic fibrosis. J Histochem Cytochem 48, 831–837. 5. Gentzsch, M., and Riordan, J. R. (2001) Localization of sequences within the C-terminal domain of the cystic fibrosis transmembrane conductance regulator which

CFTR Localization impact maturation and stability. J Biol Chem 276, 1291–1298. 6. Gentzsch, M., Chang, X. B., Cui, L., Wu, Y., Ozols, V. V., Choudhury, A., et al. (2004) Endocytic trafficking routes of wild type and DeltaF508 cystic fibrosis transmembrane conductance regulator. Mol Biol Cell 15, 2684–2696. 7. Cholon, D. M., O’Neal, W. K., Randell, S. H., Riordan, J. R., and Gentzsch, M. (2010) Modulation of endocytic trafficking and apical stability of CFTR in primary human airway epithelial cultures. Am J Physiol Lung Cell Mol Physiol 298, L304–L314. 8. Fulcher, M. L., Gabriel, S., Burns, K. A., Yankaskas, J. R., and Randell, S. H. (2004) Well-differentiated human airway epithelial cell cultures. Methods Mol Med 107, 183–206.

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9. Gentzsch, M., Cui, L., Mengos, A., Chang, X. B., Chen, J. H., and Riordan, J. R. (2003) The PDZ-binding chloride channel ClC-3B localizes to the Golgi and associates with cystic fibrosis transmembrane conductance regulatorinteracting PDZ proteins. J Biol Chem 278, 6440–6449. 10. Engelhardt, J. F., Yankaskas, J. R., Ernst, S. A., Yang, Y., Marino, C. R., Boucher, R. C., et al. (1992) Submucosal glands are the predominant site of CFTR expression in human bronchus. Nat Genet 2, 240–247. 11. Kälin, N., Claass, A., Sommer, M., Puchelle, E., and Tümmler, B. (1999) DeltaF508 CFTR protein expression in tissues from patients with cystic fibrosis. J Clin Invest 103, 1379–1389. 12. Pawley, J. B. (2006) Handbook of Biological Confocal Microscopy. Springer, New York, NY.

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Chapter 3 CFTR Regulation of Epithelial Sodium Channel Yawar J. Qadri, Estelle Cormet-Boyaka, Dale J. Benos, and Bakhrom K. Berdiev Abstract Cystic fibrosis (CF) is a lethal genetic disorder, characterized by both clinical and genetic complexities, and arises as a result of mutations in the cystic fibrosis transmembrane conductance regulator (CFTR) gene. The gene encodes a Cl− channel belonging to the ABC (ATP Binding Cassette) superfamily of transporters. The members of this superfamily use ATP hydrolysis to fulfill their function as active transporters. So far, CFTR is the only member of this family to function as a cAMP-activated Cl− channel. Intense research following the cloning of the CFTR gene has extended the role of the CFTR beyond that of a Cl− channel. One of the best recognized, yet still controversial, functions of the CFTR is its ability to modulate the functioning of other transporters. The modulation of epithelial Na+ channel (ENaC) function serves as a prime example of regulatory function of the CFTR. In this chapter, we will briefly describe an integrated protocol consisting of biochemical and electrophysiological approaches to study the regulation of ENaC by CFTR. Key words: CFTR, ENaC, bilayers, two-electrode voltage clamp, co-immunoprecipitation.

1. Introduction The cystic fibrosis transmembrane conductance regulator not only functions as an ATP- and PKA-dependent Cl− channel but also has the unique ability to influence the function of other transporters (1, 2). The modulation of epithelial Na+ channel (ENaC) is one of the best recognized, yet still debated regulatory functions of the CFTR. In this chapter, we will describe a protocol that combines biochemical and electrophysiological approaches to study the regulatory link between ENaC and CFTR.

M.D. Amaral, K. Kunzelmann (eds.), Cystic Fibrosis, Methods in Molecular Biology 742, DOI 10.1007/978-1-61779-120-8_3, © Springer Science+Business Media, LLC 2011

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2. Materials 2.1. Xenopus laevis Oocyte Preparation and Injection with cRNAs and Preparation of Oocyte Plasma Membrane Vesicles

1. Female X. laevis (Xenopus Express, Brooksville, FL, USA, or Le Bourg, France). 2. General anesthesia for amphibians. 3. Ca2+ -free OR-2 solution: 82.5 mM NaCl, 2.4 mM KCl, 1.8 mM MgCl2 , 5.0 mM HEPES (pH 7.4). 4. Collagenase type 1A. 5. Leibovitz (L-15) buffer (Sigma, St. Louis, MO). 6. cRNAs of the CFTR and ENaC in nuclease-free H2 O (see Note 1). 7. Rinsing buffer (high-K+ ): 400 mM KCl, 5 mM piperazineN,N-bis(2-ethanesulfonic acid), pH 6.8. 8. Protease inhibitor cocktail: phenylmethylsulfonyl fluoride, pepstatin, aprotinin, leupeptin. 9. DNase I. 10. Sucrose. 11. Resuspension buffer: 100 mM KCl, 5 mM MOPS, pH 6.8.

2.2. Planar Lipid Bilayers

1. The bilayer setup (Warner Instruments, Inc., Hamden, CT; see Note 2). 2. Ag/AgCl electrodes embedded in 3 M KCl/3% agar bridges (see Note 3). 3. The bathing solutions: 100 mM NaCl, pH 7.4 (see Note 4). 4. Bilayer-forming lipid: 10 mg/mL diphytanoyl phosphatidylethanolamine (Avanti Polar Lipids, Alabaster, AL) stock in chloroform. 5. Bilayer-forming lipid: 10 mg/mL diphytanoyl phosphatidylserine (Avanti Polar Lipids, Alabaster, AL) stock in chloroform. 6. Nitrogen gas. 7. Bilayer-forming solution (n-decane or n-octane). 8. Protein kinase A and MgATP (to phosphorylate CFTR Cl− channel), CFTR inhibitors (diphenylamine-2-carboxylate, glibenclamide, or anti-CFTR505-511 antibody), and ENaC inhibitors (amiloride or benzamil).

2.3. Two-Electrode Voltage Clamp

1. Two-electrode voltage clamp system including an amplifier, micromanipulators for electrode positioning, a vibration isolation table, and a Faraday cage (Warner Instruments, Inc., Hamden, CT; see Note 5).

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2. Perfusion system and recording chamber (Warner Instruments, Inc., Hamden, CT; see Note 6). 3. Electrode puller and glass. 4. Electrode solution, 3 M KCl. 5. Bath solution, primarily ND96: 96 mM NaCl, 1 mM MgCl2 , 1.8 mM CaCl2 , 2 mM KCl, and 5 mM HEPES (pH 7.4). 6. Amiloride, (IBMX). 2.4. Cell Culture, Transient Transfection

forskolin,

and

3-isobutyl-1-methylxanthine

1. Lysis buffer: PBS 1×/0.2% Triton X-100 containing R protease inhibitor cocktail (Roche Biochemical, Complete Mannheim, Germany). Cell culture solutions (see Note 7). 2. 4–15% gradient gel (Bio-Rad). 3. Running buffer (10×): 1×: 25 mM Trizma base, 192 mM glycine, 0.1% SDS. Store at room temperature. 4. Transfer buffer: 25 mM Trizma base, 192 mM glycine. Store at 4◦ C. 5. Tris-buffered saline (TBS): 150 mM NaCl, 25 mM Tris. Adjust pH to 7.5 with HCl. 6. Blocking buffer: TBS with 0.1% Tween-20 and 5% nonfat dry milk. 7. Primary and secondary antibodies diluted in antibody dilution buffer (TBS with 0.4% Tween-20, 8% glycerol, and 4% nonfat dry milk) (see Note 8). 8. 5× SDS loading sample buffer: 250 mM Tris–HCl (pH 6.8), 10% SDS, 30% glycerol, 5% β-mercaptoethanol, and 0.02% bromophenol blue.

3. Methods 3.1. Preparation and Injection of Xenopus Oocytes

1. Surgically remove oocytes from an anesthetized (ice/tricaine solution) adult female X. laevis kept at 18◦ C in chlorine-free water (3, 4). 2. Oocyte defolliculation: 2-h treatment with collagenase type 1A (1 mg/mL) in Ca2+ -free OR-2 solution. Exchange solution at least once, preferably twice. 3. Choose and isolate stage V/VI oocytes. 4. Allow oocytes to recover overnight in half-strength Leibovitz L-15 buffer.

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5. Divide oocytes into three groups (30–40 in each): ENaCs alone, ENaCs plus CFTR, and control. 6. ENaCs without CFTR group: inject oocytes with 2 ng/subunit cRNA (in nuclease-free H2 O) for each ENaC subunit (ENaC is comprised of three subunits giving a total of 6 ng cRNA; see Note 9). ENaCs with CFTR group: add CFTR cRNA (a total of 8 ng cRNA). Control group: inject oocytes with 50 nL of water only (no RNA). 7. Incubate injected oocytes for 48–72 h in L-15 or similar media. Change medium daily as some oocytes may not survive the injection procedure. 3.2. Preparation of Xenopus Oocyte Membrane Vesicles

1. Rinse injected oocytes with high-K+ buffer supplemented with 100 μM phenylmethylsulfonyl fluoride, 1 μM pepstatin, 1 μg/mL aprotinin, 1 μg/mL leupeptin, 1 μg/mL DNase I, and 300 mM sucrose (5). 2. Use a ground glass tissue grinder to homogenize oocytes in 300 μL (∼10 μL/oocyte) of the high-K+ buffer for 5 min. 3. On a discontinuous sucrose gradient (3 mL of 50% on bottom and 3 mL of 20% on the top in high-K+ buffer in the presence of protease inhibitors) layer homogenate. 4. Centrifuge at 23,500×g for 30 min. 5. Discard the top layer. 6. Collect the interface (white cloudy layer) and dilute threefold with high-K+ buffer. 7. Repeat centrifugation at 23,500×g for 30 min. 8. Discard the supernatant and resuspend pellet in 100 μL of resuspension buffer (100 mM KCl, 5 mM MOPS, pH 6.8). 9. Separate membrane vesicles into 15-μL fractions and store at −80◦ C until use.

3.3. Planar Lipid Bilayer System

1. The bilayer chamber has two parts (with cis and trans designation) (3, 4). These compartments are separated by a polycarbonate or a Teflon septum containing a small (150– 200 μm) hole. The voltage source is connected to the cis compartment of the bilayer chamber (via Ag/AgCl electrode and 3 M KCl/3% agar bridge). The current-to-voltage converter is connected to the trans compartment of the chamber (via Ag/AgCl electrode and 3 M KCl/3% agar bridge) and serves as a virtual ground (see Note 10). 2. To increase the chances of vesicle fusion, a mix of negatively charged (phosphatidylserine) and neutral (phosphatidylethanolamine) lipids is recommended. The ratio of the lipids can be varied (see Note 11). Lipids are available

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from Avanti Polar Lipids (Alabaster, AL, USA). The lipids should be stored at –20◦ C. The yellow color of the lipids is indicative of oxidation and such lipids should not be used. The solution for forming bilayer should be made daily. 3. The “painting” approach (6, 7) for bilayer formation is recommended; a bilayer is formed by applying a small amount of bilayer-forming solution over the hole in the septum that separates the cis and trans compartments. A membrane with the capacitance of 200–300 pF (0.67–0.95 μF/cm2 ) can be used for vesicle fusion. 3.4. Bilayer Incorporation

1. In a glass vial, mix 20 μL stock of diphytanoyl phosphatidylethanolamine with 10 μL stock of diphytanoyl phosphatidylserine. 2. Dry the lipid mixture under flowing nitrogen. 3. Use 35–60 μL of n-decane (or n-octane) to dissolve dried lipids (to a final lipid concentration of 12.5–25 mg/mL). 4. Add bathing solution to the cis and trans compartments of the bilayer chamber. 5. Connect the trans chamber of the bilayer system to the current–voltage converter using an Ag–AgCl electrode and 3 M KCl–3% agar. 6. Connect the cis chamber of the bilayer system to the voltage source using an Ag–AgCl electrode and 3 M KCl–3% agar. 7. Use “painting” method to form bilayer over the septum aperture using the lipid containing membrane-forming solution in n-decane described in Step 3 (see Note 12). 8. Establish the bilayer membrane formation by following membrane capacitance. 9. Place a small aliquot of the oocyte vesicle suspension into the trans chamber and wait for fusion to occur (see Note 13). 10. The voltage is changed and currents are recorded, stored, and analyzed using a computer running pCLAMP software (Axon Instruments, Burlingame, CA, USA). 11. Sensitivity to amiloride and benzamil is used to dissect ENaC activity in bilayers (see Note 14). 12. Phosphorylation by protein kinase A and MgATP is a requirement to detect CFTR Cl– channel. Also, the CFTR inhibitors (diphenylamine-2-carboxylate, glibenclamide, or anti-CFTR505-511 antibody) can be used to further identify CFTR (see Note 14).

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3.5. Two-Electrode Voltage Clamp System

1. A two-electrode voltage clamp system is designed for recordings of relatively large macroscopic currents from a single large cell such as the Xenopus oocyte in a much simpler manner as compared to the patch clamp technique. Depending on the length of the experimental protocol, tens to hundreds of oocytes can be screened in a week by a skilled operator. 2. Two sharp electrodes are used to impale a single cell. The electrodes act as either a “voltage” electrode or a “current” electrode, the former recording the transmembrane potential and the latter injecting current into the cell. Separate bath electrodes should be used to clamp the bath solution to ground, adding two Ag/AgCl wires which sit either directly in the solution or in an adjacent chamber connected to the oocyte chamber with 3 M KCl/3% agar salt bridges. This reduces the noise levels and helps remove series resistance. The agar salt bridges are advised as silver ions can be toxic to cellular processes and also the Ag/AgCl wires are sensitive to differences in chloride concentrations in the solutions. 3. Unlike single-channel bilayer recordings, two-electrode voltage clamp recordings are macroscopic measurements of all the channels expressed at the membrane of the oocyte. Though single-channel properties may be inferred, the recordings are of a large population of channels which may or may not change depending on trafficking or signaling events. This makes the technique dependent on the complexities of protein networks.

3.6. Two-Electrode Voltage Clamp Recordings of ENaC

1. The tips of two glass electrodes filled with 3 M KCl are lowered into the bath chamber and submerged in the bath solution (see Note 15). The electrical resistance of the individual electrodes should measure between 0.5 and 2 M. The resistance of the current electrode should ideally be less than that of the voltage electrode. Solution should not be actively flowing out of either electrode. 2. A single oocyte is placed within the chamber and the two electrode tips are pressed up against the oocyte. It may help to place a plastic mesh or some sort of support to keep the oocyte from moving into the chamber, though care must be taken to minimize disruption of the solution flow. The oocyte may then be impaled by the electrodes by gently advancing them until they pop through the membrane or by using the “buzz” feature on some amplifies. Minimal force is required to avoid damaging the oocyte and leaking cytosolic contents. Entry can be monitored by examining the voltage recording from the electrodes, as the uninjected oocyte

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resting membrane potential is expected to be in the –30 to –60 mV range, though large variations can be observed. 3. Although electrophysiological protocols vary, for ENaC and CFTR, the oocyte is generally held at a constant voltage, while the whole-cell current is recorded. For ENaC, –60 mV is commonly used to record the inward sodium current. Using voltage steps, it is possible to calculate a chord conductance (g) using Ohms law, Iion = gion (Vm –Vion ), where Vm is the holding potential, while Vion is the reversal potential for the ion. 4. Bath solutions are continuously applied, generally using a gravity-driven perfusion apparatus to bathe the oocyte, maintaining a constant volume of solution in the chamber. To measure the ENaC currents, solutions containing channel blockers such as 10 μM amiloride are bathed over the oocyte and currents are recorded. ENaC inhibition by amiloride should occur rapidly, though the timescale will depend on the perfusion system, and is reversible. The difference between recordings with and without amiloride can be used to determine the amiloride-sensitive ENaC currents. For CFTR, solutions containing a CFTR activator cocktail such as 10 μM forskolin with 0.2 mM 3-isobutyl-1methylxanthine (IBMX) are added. These reagents are used to increase cAMP levels within the oocyte and may take up to 10–15 min to reach full effect; therefore the current must be allowed to stabilize before measurements are made, and these effects are not rapidly reversible. The difference between recordings with and without this cocktail can be attributed to CFTR. When ENaC and CFTR are coexpressed, amiloride-sensitive currents can be measured before activation of CFTR and then repeated after activation of CFTR. The effect of these reagents on water-injected oocytes should be used as a control for endogenous currents. This can allow the isolation of CFTR or ENaC currents in the absence or the presence of the other. 5. In oocytes expressing CFTR alone, currents are not sensitive to 10 μM amiloride before or after activation of CFTR. Activation of CFTR by forskolin and IBMX leads to an increased outward chloride flux. 6. In oocytes expressing ENaC alone, application of 10 μM amiloride leads to a rapid reduction of the macroscopic current. Application of forskolin and IBMX has little to no effect on the amiloride-sensitive current. 7. In oocytes expressing both ENaC and CFTR, a smaller amiloride-sensitive current is observed before CFTR activation, showing a decrease in ENaC activity or expression.

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After amiloride washout, a forskolin- and IBMX-induced current is observed that is larger than when CFTR is expressed alone, indicating an increase in the CFTR activity or expression by the presence of ENaC. Further application of amiloride following CFTR activation shows a further reduced amiloride-sensitive current. 8. Succinctly, the presence of ENaC increases CFTR currents, while the presence of CFTR decreases ENaC currents, and activation of CFTR further reduces ENaC currents (see Note 16). 3.7. Cell Culture, Transient Transfection, Coimmunoprecipitation

1. Perform the cell culture and transient transfection under sterile technique in a laminar flow hood (8). 2. Maintain human embryonic kidney 293T (HEK293T) cells in DMEM media (GIBCO) supplemented with 10% FBS (HyClone) and penicillin/streptomycin (GIBCO) in tissue culture-treated flasks at 37◦ C with 5% CO2 . 3. One day before transfection, subculture the cells using trypsin and seed the cells in 35-mm dishes or six-well plates (see Note 17). 4. When the cells reach 80% confluence (usually on the second day), transiently transfect them with construct(s) of interest using Lipofectamine 2000 (Invitrogen). 5. Dilute 2.5 μL of Lipofectamine 2000 (2.5 μL Lipofectamine 2000/μL of cDNA) reagent with 100 μL of OptiR I and incubate at room temperature for 5 min. MEM Separately, dilute 1 μg of each cDNA construct with 100 R I. Incubate for 5 min; combine the μL of Opti-MEM diluted cDNA constructs with diluted Lipofectamine 2000 and allow 20 min for complex formation. 6. After incubation, add the transfection solution (containing the cDNA/Lipofectamine 2000 complex) to the cells in R I. Opti-MEM 7. Incubate the cells for 5 h at 37◦ C in a CO2 incubator. 8. After incubation, change the transfection solution to regular growth medium without antibiotics. 9. Perform co-IP studies after 24–48 h of incubation to allow for protein expression.

3.8. Preparation of the Samples for Detection by Western Blotting

1. Lyse the cells by adding 200 μL (per well of a six-well plate) of lysis buffer and leave on ice for 10 min. Then pipette up and down to homogenize and transfer to a centrifuge tube. 2. Discard non-soluble material with centrifugation (15,800×g for 10 min at 4◦ C).

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3. Keep 20 μL of supernatant (lysate) and incubate the rest of the supernatant with 1 μg of carboxy-terminal CFTR monoclonal antibody (R&D Systems, 24-1) cross-linked to 25 μL A/G agarose beads (Santa Cruz Biotechnology) for 2 h at 4◦ C (see Note 18). 4. Pellet the beads with centrifugation (3,000×g for 2 min) and wash three to five times with cold PBS containing 0.2% Triton X-100. 5. Elute the proteins from the beads by adding 5 μL of 5× SDS sample loading buffer and incubate for 10 min at 37◦ C. 3.9. SDS-PAGE and Transfer

1. When using precast ready gels, remove the comb and rinse the wells with distilled water. 2. Remove the seal located at the bottom of the gel as indicated in the manufacturer’s instructions. 3. Insert the gel in the unit and place them in the tank (mini PROTEAN system from Bio-Rad). 4. Add the diluted running buffer to the tank. 5. Load each supernatant from the co-IP to the wells. Include one well that will contain the prestained molecular weight markers (Precision Plus from Bio-Rad). 6. Complete the assembly of the unit and connect to a power supply that will be set to 200 V for 30–45 min. The gel can be run at room temperature or at 4◦ C. 7. Set up the transfer by immersing the PVDF membranes into methanol for 5–10 min. The PVDF membrane should be of the size of the gel. 8. Equilibrate the membranes by immersing them in transfer buffer for 15 min. 9. Take the gels (containing the proteins) and incubate in transfer buffer for 10 min. 10. Assemble the gel and the PVDF membrane so that the separated proteins present in the gel will be transferred to the PVDF membrane. The orientation of the gel can be marked by cutting a corner of the membrane. 11. Set up the assembly into a Bio-Rad Trans Blot Cell and add the cold transfer buffer (see Note 19). Connect to a power supply that will be set to 60 V for 1.5 h (if Trans Blot Cell with wire electrodes) or 80 V for 35 min (if Trans Blot Cell with plate electrodes). The transfer unit should be placed at 4◦ C.

3.10. Western Blotting for CFTR and ENaC

1. Unassemble the gel/PVDF membrane sandwich and rinse the membrane with distilled water. Make sure to place the side of the PVDF membrane that was in contact with the

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gel face up. This is the side that will be exposed to the film in Step 10. The molecular weight markers should now be present on the membrane. 2. Use a container that is close to the size of the PVDF membrane (blot). Add enough blocking solution to cover the blot (≈10 mL) and leave on rotator for 1 h at room temperature. 3. Add the anti-ENaC Ab that recognizes the different ENaC subunits diluted in antibody dilution buffer. Leave for 2 h at room temperature or overnight at 4◦ C. 4. Remove the primary antibody and wash with TBS/0.1% Tween-20 by adding 10 mL five times for 5 min each. The box containing the blot is placed on a rotator to enhance the washing. 5. Incubate the blot with 10 mL blocking solution for 10 min at room temperature. 6. Add the secondary antibody (anti-rabbit HRP from Pierce) at a dilution of 1/10,000 for 1 h at room temperature. 7. Repeat Step 4 (washings). 8. Remove the blot from the box and blot it to remove excess washing buffer by blotting the membrane with Kim-Wipes. Place in a clean box and add West Pico (Pierce) solution. Leave the solution for 5 min at room temperature. 9. Take the blot and remove the excess West Pico solution by blotting the membrane with Kim-Wipes. 10. Place the blot in a sheet protector and place in an X-ray film cassette. 11. In a dark room, add a film on the top of the sheet protector containing the blot and expose the film for a few seconds to several minutes depending on the intensity of the signals (see Fig. 3.2 for an example of the results). 12. Always run controls to assure the specificity of the association (see Note 20).

4. Notes 1. It is important that only nuclease-free water and appropriately prepared (nuclease-free) material are used to prevent cRNA degradation. 2. The bilayer setup can be assembled from both manufactured and homemade components (9–11). On the other hand, the complete bilayer setup (vibration isolation table,

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Faraday cage, amplifier, Bessel filter, cups and chambers, and various accessories) can be purchased from Warner Instruments. 3. Ag/AgCl electrodes are prepared by “chloriding” a silver wire in bleach for 10 min. 4. The content of the bathing solutions is dependent on the channel of interest. 5. The two-electrode system is contained within a Faraday cage to remove external electrical interference and requires the use of a stable platform to minimize vibration within the system. Although a fully active vibration isolation table is recommended by some, it is not essential. Furthermore, the micromanipulators used do not need to be electronically controlled or as finely controlled in movement as those required for patch clamp system as the dimensions traveled are much larger with the oocyte. However, drift or uncontrolled movements are dangerous, especially with long experiments where irreversible mechanical damage can occur to the oocytes. 6. There are multiple chambers available. Homemade chambers can be designed using a simple Petri dish. It is important that the perfusion system and the chamber used allow for relatively stable levels of solution with relatively laminar flow as the oocytes are sensitive to mechanical damage and the electrical recordings are sensitive to solution levels. 7. Prewarm the DMEM, Opti-MEM, and trypsin–EDTA solution to 37◦ C in a water bath; before transferring to the hood, decontaminate by wiping the surface of the bottles/tubes with 70% ethanol. 8. The addition of sodium azide to a final concentration of 0.02% can save primary antibodies for subsequent experiments. 9. cRNA for oocyte injection is made and purified with the Ribomax kit plus RNA cap analogue and GeneClean kits, respectively. 10. A magnetic bar and a stirring device, along with a perfusion system, are components of a bilayer setup. The bathing solutions should be made daily and filtered. The experiments are usually performed at 25 ± 10◦ C. 11. The lipid types/ratios as a bilayer-forming solution can be varied. 12. The pre-treatment of septum aperture with bilayer-forming solution (3–5 min before filling the compartments of chamber with bathing solution) helps bilayer formation and stability.

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13. The fusion can be promoted by bringing a fire-polished glass rod dipped into oocyte vesicle suspension in close proximity to the pre-formed planar bilayer membrane from the trans side. If this approach is implemented, the bilayer membrane should be clamped at a negative voltage. Following incorporation of ENaC into the planar bilayer, channels of uniform conductance with well-defined gating transitions become apparent. 14. The fusion of ENaC-containing vesicles results in the appearance of the conductance with well-defined gating. The CFTR is known to decrease the open probability (the time channel spends in open time) of ENaC. The CFTR effect on ENaC can be assessed using vesicles containing CFTR and ENaC. Alternatively, the ENaC vesicles can be incorporated first into bilayers followed by coincorporation of the CFTR-containing vesicles. However, the co-incorporation approach is technically difficult, and we recommend the incorporation of the vesicles containing both CFTR and ENaC. First, the vesicles are incorporated into bilayers and ENaC activity is measured. Then, CFTR activity is elicited by phosphorylation of the channel. Further, the CFTR identity is dissected by applying glibenclamide (the CFTR inhibitor), while the ENaC activity is blocked by amiloride (Fig. 3.1). 15. The glass electrodes tend to get clogged after extended use but they may be reused between oocytes if the resistance stays within 0.5–2.0 M. Multiple electrodes can be prepared in advance using a large container with clay to hold the glass in place and keep the tips from breaking. Depending on conditions, they may need to be covered to prevent dust and debris from accumulating, but these tips are less sensitive than patch electrodes. As with most glass electrodes backfilled with electrolyte solutions, bubbles are a serious problem, but these tips are generally easier to fill than patch clamp electrodes and can be easily backfilled with a MicroFil 34-gauge needle from World Precision Instruments, Inc. The 3 M KCl can be pushed through a syringe filter to remove debris. 16. Studies of ENaC and CFTR in the oocyte system are conflicting (see Ref. 2). In our hands, the results we share are readily apparent. The work of Georg Nagel has suggested that there is no electrophysiological interaction, finding no basal inhibition of ENaC by the presence of CFTR and no inhibition following activation of CFTR (12, 13). These authors suggest that the observations of an interaction are merely artifacts due to a combination of CFTR’s large conductance and a high series resistance due to improperly

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Fig. 3.1. CFTR modulation of αβγ-ENaC in bilayers. First, oocyte vesicles expressing ENaC alone were fused with bilayers (top trace) which spent roughly 52% of time in open state. The time spent in open state by ENaC decreased to roughly 37% with subsequent fusion of the oocyte vesicles expressing CFTR into bilayers (second trace; also, the same effect was observed with the bilayer fusion of the oocyte vesicles co-expressing both CFTR and ENaC). The phosphorylation (PKA+MgATP, third trace) revealed the presence of the CFTR which was inhibited by anti-CFTR antibodies (fourth trace, left panel) leaving only ENaC activity which in turn was inhibited by amiloride (fifth trace, left). If, on the other hand, amiloride was applied after phosphorylation, the CFTR activity was evident (fourth trace, right panel), which in turn was inhibited by DPC addition (fifth trace, right panel). This research (15) was originally published in The Journal of Biological Chemistry. Ismailov et al., 1996; 271:4725–4732. © The American Society for Biochemistry and Molecular Biology.

configured or absent bath electrodes (12, 13). Others have found no basal inhibition of ENaC by CFTR but rather one dependent solely upon the Cl– channel activity of CFTR (for review see 14). The discrepancies between the results have yet to be satisfactorily resolved. 17. The number of cells seeded should be that they reach 80% confluence the next day.

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Fig. 3.2. (a) β-ENaC co-immunoprecipitation with CFTR. Only simultaneous presence of both CFTR and β-ENaC, with the combinations of α/β-ENaC, β/γ-ENaC, or β-ENaC with CFTR (lanes 2–5), resulted in co-IP signal. The absence of either CFTR (lane 1) or β-ENaC (lane 6) eliminated the co-immunoprecipitation signal. The presence (lane 7) or the absence (lane 8) of β-ENaC signal was confirmed in lysates. The bottom panel of the figure shows the same blot reprobed with anti-CFTR Ab to confirm the presence of CFTR in the immunoprecipitate. (b) β-ENaC does not immunoprecipitate with the Cl– channel ECFP–CLCN1. Anti-GFP monoclonal Ab JL-8 (BD Living Colors) was used to immunoprecipitate CLCN1, and the blot was probed with the β-ENaC Ab. The expression of CLCN1 in the cells was confirmed using GFP Ab (lanes 2 and 4). An Ab against β-ENaC failed to detect a co-IP signal in immunoprecipitate of cells co-expressing β-ENaC and CLCN1 (lane 4, bottom panel). The expression of β-ENaC in the lysates was confirmed (lanes 1 and 2). This research (8) was originally published in The Journal of Biological Chemistry. Berdiev et al., 2007; 282:36481–36488. © The American Society for Biochemistry and Molecular Biology.

18. Keep 20 μL of cell lysate to confirm the expression of the ENaC subunits (see Fig. 3.2a, last two lanes). 19. Assemble the gel/PVDF membrane like a sandwich. Add the first sponge that was soaked in transfer buffer on the top of the black plate, then add a piece of Whatman paper that was wetted in transfer buffer, add the gel, then the PVDF membrane, and another piece of Whatman paper (wetted

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in transfer buffer). At this step, use a roller to eliminate any bubble that could be present between the gel and the PVDF membrane. Add the second sponge and close the sandwich with the red plate. 20. To check for the possibility that ENaC subunits bind nonspecifically to the beads, the ENaC subunits are expressed and incubated with non-specific antibody cross-linked to A/G beads. It is also possible that CFTR and ENaC binding occurs postlysis. Therefore, two sets of cells are transfected with ENaC subunits or CFTR cDNA. Mix these two lysates and perform coIP experiments; no ENaC signal suggests that the interaction between CFTR and ENaC does not occur post-lysis. Additionally, always reprobe the same blot with anti-CFTR Ab to confirm the presence of CFTR in the IP (Fig. 3.2a, bottom panel). Finally, substitute the CFTR with another Cl– channel that is not thought to interact with ENaC (ECFP–CLCN1, for example) and perform co-IP experiments. The absence of the co-IP signal suggests that ENaC does not interact with the chloride channel CLCN1 (Fig. 3.2b); this strengthens the specificity of the CFTR and ENaC association. As can be seen from Fig. 3.2b, we were unable to co-IP β-ENaC with ECFP–CLCN1. In this approach we used anti-GFP monoclonal Ab JL-8 (BD Living Colors) that recognizes CFP protein to IP CLCN1 and then probed with the β-ENaC Ab. The expression of CLCN1 in the cells was confirmed using GFP Ab (Fig. 3.2b). The cells expressing αβγ-ENaC and/or ECFP–CLCN1 were IPed with an Ab against GFP. While this Ab detected ECFP–CLCN1 in both crude lysates and IP of ECFP–CLCN1-expressing cells, an Ab against β-ENaC failed to detect a band in IP of co-expressing cells (top panel and right-hand lanes of bottom panel). However, this Ab did detect βENaC in the crude lysates from cells expressing either αβγ-ENaC alone or those co-expressing ECFP–CLCN1. These data suggest that β-ENaC does not interact with the chloride channel CLCN1.

Acknowledgments ENaC cDNAs were a kind gift of Dr B. Rossier (University of Lausanne, Lausanne, Switzerland). This work was supported by NHLBI grant R21HL085112 (BKB), UAB Health Services Foundation General Endowment Fund (BKB), and NIH grant DK37206 (DJB).

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References 1. Kunzelmann, K. (2001) CFTR: interacting with everything. News Physiol Sci 16, 167– 170. 2. Berdiev, B. K., Qadri, Y. J., and Benos, D. J. (2009) Assessment of the CFTR and ENaC association. Mol Biosyst 5, 123–127. 3. Awayda, M. S., Ismailov, I. I., Berdiev, B. K., and Benos, D. J. (1995) A cloned renal epithelial Na+ channel protein displays stretch activation in planar lipid bilayers. Am J Physiol 268, C1450–C1459. 4. Ismailov, I. I., Awayda, M. S., Berdiev, B. K., Bubien, J. K., Lucas, J. E., Fuller, C. M., et al. (1996) Triple-barrel organization of ENaC, a cloned epithelial Na+ channel. J Biol Chem 271, 807–816. 5. Perez, G., Lagrutta, A., Adelman, J. P., and Toro, L. (1994) Reconstitution of expressed KCa channels from Xenopus oocytes to lipid bilayers. Biophys J 66, 1022–1027. 6. Mueller, P., Rudin, D. O., Tien, H. T., and Wescott, W. C. (1962) Reconstitution of cell membrane structure in vitro and its transformation into an excitable system. Nature 194, 979–980. 7. Mueller, P., Rudin, D. O., Tien, H. T., and Wescott, W. C. (1962) Reconstitution of excitable cell membrane structure in vitro. Circulation 26, 1167–1171. 8. Berdiev, B. K., Cormet-Boyaka, E., Tousson, A., Qadri, Y. J., Oosterveld-Hut, H. M. J., Hong, J. S., et al. (2007) Molecular proximity of cystic fibrosis transmembrane conductance regulator and epithelial sodium channel assessed by fluorescence resonance

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energy transfer. J Biol Chem 282, 36481– 36488. Alvarez, O. (1986) How to set up a bilayer system, in (Miller, C., ed.), Ion Channel Reconstitution. Plenum Press, New York, NY, pp. 115–139. Alvarez, O., Benos, D. J., and Latorre, R. (1985) The study of ion channels in planar lipid bilayer membranes. J Electrophys Tech 12, 159–177. Hanke, W., and Schlue, W. R. (1993) Planar Lipid Bilayers. Academic, San Diego, CA. Nagel, G., Szellas, T., Riordan, J. R., Friedrich, T., and Hartung, K. (2001) Nonspecific activation of the epithelial sodium channel by the CFTR chloride channel. EMBO Rep 2, 249–254. Nagel, G., Barbry, P., Chabot, H., Brochiero, E., Hartung, K., and Grygorczyk, R. (2005) CFTR fails to inhibit the epithelial sodium channel ENaC expressed in Xenopus laevis oocytes. J Physiol 564, 671–682. Kunzelmann, K. (2003) Control of membrane transport by the cystic fibrosis transmembrane conductance regulator, in (Kirk, K. L., and Dawson, D. C., eds.), The Cystic Fibrosis Transmembrane Conductance Regulator. Kluwer Academic/Plenum Publishers, New York, NY, pp. 55–93. Ismailov, I. I., Awayda, M. S., Jovov, B., Berdiev, B. K., Fuller, C. M., Dedman, J. R., et al. (1996) Regulation of epithelial sodium channels by cystic fibrosis transmembrane conductance regulator. J Biol Chem 271, 4725–4732.

Chapter 4 Methods for Evaluating Inflammation in Cystic Fibrosis Assem G. Ziady and Pamela B. Davis Abstract Cystic fibrosis is characterized by excessive pulmonary inflammation, which presents early in life and becomes self-sustaining, eventually leading to the destruction of the lung. Treating inflammation is one of the most pressing needs in CF therapy and has been shown to slow lung function deterioration. However, it remains unclear whether excessive inflammation is a direct result of CFTR dysfunction, and thus innate, or develops in response to early stimulation of inflammatory pathways. Here, we will discuss clinically relevant studies and the methods employed by them. We will focus on investigations in cell and animal models as well as patients. Our discussion will describe the character of pulmonary inflammation in CF and present potential therapeutic approaches that can ameliorate excessive responses and improve disease prognosis. Key words: Inflammation, cytokines, transcription factors, anti-inflammatory agents.

1. Introduction Inflammation is a hallmark of cystic fibrosis (CF). Early in life, patients begin to exhibit exaggerated inflammatory profiles, especially in the lungs (1–4) in response to infection. Over time, these responses become more persistent and contribute to the destruction of lung tissue, pulmonary function deterioration, respiratory failure, and death. It is unclear whether infection causes the excessive inflammation observed in CF or whether dysfunction of the cystic fibrosis transmembrane conductance regulator (CFTR) in itself results in aberrant cell signaling that gives rise to abnormal inflammatory responses. A number of studies have described increases in pathways that promote inflammation (1–10) and decreases in pathways that are anti-inflammatory M.D. Amaral, K. Kunzelmann (eds.), Cystic Fibrosis, Methods in Molecular Biology 742, DOI 10.1007/978-1-61779-120-8_4, © Springer Science+Business Media, LLC 2011

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(11, 12), even in the absence of bacteria. However, many of these studies were conducted in immortalized cell line models of CF and thus require validation in animal models or primary CF patient tissues. Clinically, this has been a difficult issue to address. Reports of increased inflammatory cytokine production in the healthy non-infected lungs of CF newborns (13) do not preclude the possibility of recent infections that had cleared by the time of examination. Therefore, it is difficult to distinguish between the presence of innate abnormalities in inflammatory signaling and residual inflammation from prior infection. Other reports have focused on reduced airway clearance, which is directly linked to the lack of chloride ion secretion through functional CFTR and exaggerated sodium absorption through the epithelial sodium channel (ENaC), as the cause of increased bacterial adherence and established infections that perpetuate inflammatory signaling (14). Nevertheless, it is not disputed that inflammation exists and that controlling it provides significant benefits to patients (15, 16). Evaluations of inflammation and its impact on disease progression have been conducted on two levels: assessing inflammation using clinical samples obtained from patients and elucidating mechanisms that influence inflammation in primary cells and models of CF. Studies in patients have revealed that excessive levels of inflammatory cytokines (4–7) are produced in response to infection. Anti-inflammatory drugs, such as ibuprofen, are effective in limiting lung deterioration (15), but adverse effects have discouraged the use of both steroidal and non-steroidal anti-inflammatory drugs (16, 17). Nevertheless, alternative antiinflammatory therapies are an active area of research (18). To evaluate mechanisms that contribute to inflammation, investigators have relied on a variety of cell and animal models. The development of a number of immortalized matched epithelial cell pair models has allowed researchers to conduct the majority of mechanistic studies in the field, identifying the contribution of aberrant ion transport (14), protein expression (19) and processing (20), redox signaling (21), and lipid processing (22) to the proinflammatory phenotype. However, the field has called for the confirmation of these relationships in vivo or in primary cell culture, so methods have been developed to allow for the culture of primary airway epithelia collected from tissue discarded at transplantation, explants, or nasal scrapings from patients, as well as differentiated cells (23–25). The development of animal models has also been very active. However, no CF mouse model develops spontaneous pulmonary inflammation, and manipulations that force the establishment of infection are required to elicit differential inflammatory responses in the lung. Furthermore, there are significant anatomical differences between mice and humans; for example, submucosal glands, which are present in the human airway and

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believed to be a key site of CF airway disease, are rare in the mouse airway (26). Recent introduction of the ferret (27) and pig (28) models of CF that develop many of the phenotypes associated with CF without manipulation is very promising. Much of the current understanding of the inflammatory process in CF patients has come from sampling the airway surface liquid (ASL) via bronchoscopy and bronchoalveolar lavage (BAL). Cytokine content of BAL remains the best surrogate measure of inflammation in patients (4). BAL from young infants, even if not infected, contains elevated proinflammatory cytokines (4, 8). Studies focused on assessing inflammatory cell behavior in the CF lung find persistent infiltration of neutrophils in response to chemoattractants secreted by macrophages, epithelial cells, and the neutrophils themselves (29). Neutrophils also secrete oxidants and proteases and are the major source of DNA that increases the viscosity of CF sputum (4). In cell culture, investigators have assessed inflammation by measuring cytokines secreted by CF model cell lines or primary culture of CF epithelia following stimulation, compared with normal controls. Other studies have focused on the examination of intracellular signaling cascades including genes, proteins, or lipids that impact inflammation by gene array (30, 31), proteomic analyses (19, 21), or direct measurement. In general, these reports find an increase in the expression and/or activity of proinflammatory and a decrease in anti-inflammatory mediators. Many of these studies and their findings have been extended to and confirmed in CF animal models by sampling the BAL (32), micro-dissected (33) or whole lung (21, 32), and/or bowel tissues (34). For the most part, the proinflammatory characteristics observed in stimulated cells in cell culture have been reproduced in CF mouse tissues stimulated with inflammatory cytokines (32), bacteria (23, 30), or bacterial components (18). But the lack of inflammation in the unstimulated state continues to raise concerns about this model. Therefore, investigators have begun to evaluate cytokine, gene, and protein expression profiles in newer CF models in ferrets and pigs, both of which spontaneously develop inflammatory disease.

2. Materials For the purposes of this discussion, we will focus on airway epithelial cell models of CF. For in vivo studies, although CF models are now available in mice, ferret, and pig, we will outline materials and methods for use in studies of inflammation in the lungs of mice ∼25 g in weight. However, these methods can be extended to larger animal models.

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2.1. Cell Culture, Stimulation, and Treatment with Compounds

1. CF cell models of choice: We commonly use two airway epithelial cell lines and human primary tracheal epithelia (HTE) for our studies. For the cell lines, we use the 9HTEo− pCEP and pCEP-R cell pair and the 16HBEo− sense (S) and antisense (AS) cell pair. 9HTEo− cells are immortalized human tracheal epithelia that are stably transfected with an empty pCEP construct to produce normal controls or a pCEP-R construct that encodes for the regulatory (R) domain of CFTR to produce cells that lack CFTR function, but not CFTR itself (35). 16HBEo− are human bronchial epithelial cells that are stably transfected with a sense (S, normal control) or antisense (AS, CF cell pair) construct of nucleotides 1–131 of CFTR and which lack CFTR (35). For primary cell culture, we obtain cells from necropsy under institution-approved protocols. 2. 10% FCS in appropriate cell culture media as needed (see Note 1). For primary cell culture we use Ultra-ser G (Biosepra Inc., Marlborough, MA). 3. Selection compounds, such as hygromycin or G418, as needed. 4. Antibacterial and/or antimycotic compounds, such as penicillin, streptomycin, or fungizone, as needed (see Note 2). 5. Tissue culture-treated welled plates with or without semipermeable membrane inserts (Costar; Corning Inc., Corning, NY). 6. PBS: Dulbecco’s phosphate-buffered saline. 7. For mass spectrometric quantitation of labeled protein, protein labels (i.e., non-abundant isotopes of carbon or nitrogen) may be added to cell culture media as a supplement. 8. Ion channel modulating compounds, such as inhibitors of CFTR, which can produce the CF phenotype in treated cells (23). We usually use 20 μM CFTRinh -172 (SigmaAldrich, St. Louis, MO) in DMSO (see Note 3). 9. Stimulants such as bacteria, cytokines, or bacterial components can be used to activate inflammatory signaling. We have used 105 –109 cfu/mL of a laboratory strain of Pseudomonas aeruginosa, PAO1 (23, 32), 1 μg/mL LPS derived from P. aeruginosa (Sigma-Aldrich), or 10–100 ng/mL recombinant human tumor necrosis factor alpha (TNF-α) and 10–100 ng/mL recombinant human interleukin 1 beta (IL-1β). 10. For anti-inflammatory agents, we have used triterpinoids such as 2-cyano-3,12-dioxooleana-1,9 (11)-dien-28-oic


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acid (CDDO, 300 nM), pioglitazone (Cayman Chemical), and antioxidants such as N-acetyl cysteine. 2.2. Animal Models, Stimulation, and Treatment with Compounds

1. A number of CF animal models are available in mice (see Note 4). Some commonly used mouse models bred into the C57BL/6 background are the UNC knockout mouse (B6.129P2-Cftrtm1Unc ) and mutant murine Cftr mice such as the R117H mutant mouse (B6.129S6-Cftrtm2Mrc ). The gut-corrected knockout mouse (Cftrtm1Unc -TgN(FABPCFTR)#Jaw) is a more robust model that exhibits good survival. Ferret (27) and pig (28) models of CF are also available for study (see Note 5). 2. There is an age and sex effect on inflammatory responses. We prefer male mice that are 6–12 weeks old. 3. Animals should be housed in a sterile environment such as microisolator cages for mice. 4. Our preferred anesthetic cocktail is made up of 2.13 mg/mL xylazine, 0.36 mg/mL acepromazine, and 10.75 mg/mL ketamine. 5. One milliliter syringe with 27.5 or 30 gauge needle for intraperitoneal (IP) or intravenous (IV) injection, respectively. Larger gauge needles should be used as needed for larger animals. 6. To simplify IT administration or intubation of mice, a tilting workstation (Hallowell, Pittsfield, MA), animal laryngoscope (PennCentury, Philadelphia, PA) fitted with a magnifier to improve visualization, and a MicroSprayer (PennCentury, Philadelphia, PA) are useful. 7. For intratracheal (IT) administrations in mice, a 22 gauge plastic catheter (Abbocath; Abbot Laboratories) is used. Bronchoscopes have been used for larger animals. 8. Stimulation of inflammation is achieved with bacteria or their components. We have used 106 –109 cfu/mouse of PAO1 (23, 32) or 40 μg/kg LPS derived from P. aeruginosa (Sigma-Aldrich). 9. For anti-inflammatory agents, we have used 10–50 μg/kg CDDO, 30 mg/kg pioglitazone (Cayman Chemical), and antioxidants such as NAC. 10. A 24 or 18 gauge feeding needle can be used for mouse BAL collection or gavage, respectively. Bronchoscopes have been used for larger animals. 11. Enclosed chamber connected to a carbon dioxide gas tank and a charcoal filter (see Note 6).

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12. PBS: Dulbecco’s phosphate-buffered saline. 13. Protease inhibitor cocktail: one Complete Mini protease inhibitor tablet (Roche) + 10 mL PBS. 2.3. Measurement of Secreted Cytokines and Chemokines

1. Media collected from the apical surface of cultured cells or BAL fluid collected from animal. 2. Individual ELISA analysis kits (R&D Systems or Antigenix America Inc., for example) are widely available for the sensitive measurement of commonly measured proinflammatory mediators including IL-1β, IL-2, IL-4, IL-6, IL-8, IL-9, IL-19, GM-CSF, and TNF-α. In mice, the chemokine macrophage inflammatory protein (MIP)-2 and KC/N51 are analogs of human IL-8. ELISA kits are also available for the measurement of anti-inflammatory cytokines, such as IL-10. 3. An alternative to individual kits is the use of multi-analyses kits that measure a number of cytokines in the same biological sample. We have used the LINCOplex Multiplex kit (Linco Research, St. Charles, MO). 4. Urea detection kit for the measurement of urea in serum. A number of sensitive kits are available, such as the Infinity Urea Liquid Stable Reagent (Thermo Fisher Scientific, Inc.). 5. PBS: Dulbecco’s phosphate-buffered saline. 6. Protease inhibitor cocktail: one Complete Mini protease inhibitor tablet (Roche) + 10 mL PBS. 7. To assess bacterial load in fluids or tissue homogenates, sterile blood agar plates are used.

2.4. Histology and Immunohistochemical Staining

1. BAL or lungs harvested from CF mice and normal littermate. 2. Tissue freezing medium, such as OTC (Triangle Biomedical Sciences). 3. Paraformaldehyde dissolved in PBS at 2.5% wt/vol. 4. Paraffin wax and xylene (Thermo Fisher Scientific, Inc.). 5. Tissue microtome. 6. Hematoxylin and eosin stains (Sigma-Aldrich, St. Louis, MO). 7. Protein-of-interest specific primary antibody (Ab) and secondary HRP conjugated, or similar, Ab (see Note 7). 8. 10 mM sodium citrate buffer (pH 6.0). 9. Solution of 3% hydrogen peroxide in PBS. 10. Suitable substrate for HRP or otherwise Ab-conjugated catalytic enzyme.


2.5. One- and Two-Dimensional SDS-Polyacrylamide Gel Electrophoresis (SDS-PAGE)

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1. Rehydration/solubilization buffer: 7 M urea, 2 M thiourea, 1% DTT, 1% CHAPS, 1% ampholytes, and 1% Triton. Mix 2.1 g urea, 0.8 g thiourea, 50 mg CHAPS, 50 mg DTT, 50 μL BioLytes (BioRad), 50 μL Triton X-100. Bring volume up to 5 mL with water and dissolve with shaking (see Note 8). 2. Loading dye stock solution: 1% bromophenol blue in water. 3. Electrode wicks (Biorad), wetted with water. 4. 1D gel band or 2D gel IPG pI strip equilibration buffer: mix 5.4 g urea, 0.3 g SDS, 3.8 mL 1.5 M Tris-HCl (pH 8.8), and 3 mL glycerol in a 50 mL centrifuge tube. Adjust the total volume to 15 mL with water. Dissolve by shaking. 5. Protein reducing reagent: 120 mg DTT in 7.5 mL equilibration buffer. 6. Protein alkylation reagent: 150 mg iodoacetamide in 7.5 mL equilibration buffer with 100 μL 1% BPB. 7. Agarose for sealing 2D gel IPG pI strip (Biorad). This reagent can be stored at RT and used repeatedly over several months. 8. Running buffer suitable for purchased precast gel. For example, 1X Biorad Tris-glycine buffer. Cool on ice before use. 9. To decrease sample contamination, the use of precast gels is recommended. We use Criterion precast 1D or 2D gels (Biorad). 10. Gel-fixing solution: 50% ethanol and 10% acetic acid in water. 11. Protein label, such as GelCode BlueTM Coomassie stain (Pierce Biotechnology, Inc., Rockford, IL).

2.6. Western Blot Analyses

1. Primary Ab, suitable for Western blotting, raised against protein of interest. For inflammatory signaling, we have blotted for transcription factors such as NF-κB (18) or PPARγ (33). 2. Secondary Ab raised against the primary Ab species and conjugated to detectable enzyme, such as HRP. 3. 1D or 2D gel of CF and normal control cells or tissues. We often run different fractions of the cell (e.g., nuclear or cytoplasmic). 4. Immunoprecipitation (IP) buffer: 1% Triton X-100, 150 mM NaCl, 10 mM Tris, pH 7.4, 1 mM EDTA, and protease inhibitor cocktail. 5. Transfer buffer: 25 mM Tris, 190 mM glycine, and 10–20% methanol (vol/vol) in water. SDS

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(0.1–0.5% wt/vol) addition is sometimes necessary for the transfer of large proteins (>40 kDa). 6. Wash buffer: Tris-buffered saline (pH 7.4) with 1% Tween. 7. We use different blocking solutions depending on the protein of interest. We commonly use 5% nonfat dry milk in TBS-T or membrane blocking solution (Invitrogen, Inc.). 8. Polyvinylidene fluoride (PVDF) or nitrocellulose membranes (Millipore, Bedford, MA). 9. Enhanced chemiluminescence (ECL) reagent (Pierce, Rockford, IL). 2.7. Analysis of Promoter and/or Transcription Factor Activity

1. For evaluation of promoter activity, we use a luciferase assay-based analysis. For example, we have used firefly luciferase expression cassettes under the control of promoters for NF-κB (18), IL-8 (18), or Nrf2 (21, see Note 9). A control plasmid codes for Renilla luciferase under the control of a viral promoter (e.g., SV40). 2. For analysis of transcription factor activity, we use an ELISA-based approach such as the TransAM NF-κB p50 assay (Active Motif, Carlsbad, CA). 3. Cytoplasmic and nuclear fraction kit (Panomics). 4. Dual-Luciferase Reporter assay system (Promega, Madison, WI). 5. For examinations in cell culture, a suitable transfection reagent (see Note 10), such as LipofectinTM or Lipofectamine (Invitrogen, Inc.). 6. Plate reading luminometer. 7. For studies in animals, transgenic mice can be engineered to express luciferase under the control of a promoter of interest. 8. Animals can be imaged by a real-time bioluminescence imaging instrument, such as the Xenogen 200 (Caliper Life Sciences, Hopkinton, MA), which employs a CCD camera to sensitively detect photons emitted from tissues in living animals. 9. Small animal (7 μm) is mucus/ASL. Right: typical pH scan from the same culture 5 days later. (d) Summary data of pH taken from (c). pH in the PCL at 2 and 7 days (closed bars) and pH in the mucus layer (open bars). All n = 6. ∗ denotes p < 0.05 between PCL and mucus pH. † denotes p < 0.05 between data points at 2 vs. 7 days.

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we can record changes in pH using this method in standard solutions over the physiological range (pH 6.6–7.6). To further verify this methodology, we measured the SNARF1-dextran emission ratios in pre-washed cultures after applying 20 μL of standard Ringer solution (pH 7.4) to the mucosal surface of an airway culture. Confocal scanning 2 h after Ringer addition revealed no depth-dependent change in pH (Fig. 5.3b). However, 48 h postRinger addition, a marked gradient is clearly visible, with an emission ratio indicative of a lower pH in the region (1–7 μm) associated with the PCL and a significantly higher emission ratio in the upper limit of the mucus-containing region >7 μm (Fig. 5.3c). However, after 7 days, when the PCL had collapsed and mucus had been allowed to accumulate, the emission ratio had increased and the gradient was less pronounced. Comparing the pH values in the 7 μm periciliary region in the presence and absence of a PCL (Fig. 5.3d), it can be seen that the pH has significantly risen from 6.7 to 6.95. Similarly, the pH in the top ∼7 μm of the mucus region had also increased although in a less pronounced fashion (pH 6.95–7.1). Importantly, as the PCL became depleted of liquid and increased its mucus content, the PCL pH rose (Fig. 5.3d). To visualize the cells, they can be labeled with cell-permeant acetoxymethyl ester (AM)-based dyes. Many live cell dyes are available commercially and we have successfully used calcein-AM to stain live cells in the epithelial layer (Fig. 5.2b). A color that is different to the one used to label the ASL and/or the mucus should be picked. We have also used SNARF1-AM as a cellular dye, even though its original purpose is as a pH-sensitive dye. Furthermore, other probes undoubtedly will prove useful in the future. Mucus can be labeled using fluorescent microspheres (100 nm), which co-localize to the “strands” in the mucus layer (Fig. 5.1c) and provide a general idea of mucus location in the ASL. Mucus transport rates can be measured using larger fluorescent microspheres (∼1 μm). There are many commercially available probes and dyes which are suitable for these measurements depending on the investigators’ individual needs. Extensive lists of probes and dyes can be found online at several vendors, including Molecular Probes. We have often triple-labeled our cells. For example, the ASL is labeled with Texas Red-dextran and the mucus is labeled with 100 μM FITC beads to give an indication of ultrastructure and also with 1 μm ultraviolet beads to measure rotational mucus transport rates. 1.2. Labeling of ASL for Confocal and Epifluorescence Microscopy

To standardize ASL height measurements, the cultures are prewashed to remove excess ASL and mucus, often at a predetermined time, in order to ensure similar starting ASL volumes. This is necessary since the airway cultures are primarily Na+ /volume

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absorbing and absorb excess ASL volume with time until a steadystate height is reached (8), similar to the proximal airway epithelia (11). Thus, cultures may be washed immediately before starting an experiment to study them under mucus-free conditions or may be washed 24–48 h before the experiment to allow mucus to accumulate and to facilitate the production of a “natural” ASL. Moreover, after washing, all liquid may be aspirated, so that ASL can be tracked from minimal starting levels (25). This is especially useful to look at ASL secretion. However, to study absorption, or absorption followed by secretion, a bolus of liquid (i.e., 20 μL) may be deposited on airway surfaces and followed with time (8). The desired fluorescent probes may be added either (i) days earlier, (ii) with the final wash of the mucosal surface followed by aspiration of the ASL to minimal levels, or (iii) in a bolus of liquid (i.e., 20–50 μL) at the start of the experiment. 1.3. Use of Perfluorocarbons

Since the ASL can easily evaporate when a culture is exposed to room air, care must be taken to avoid this. One approach is to provide a highly humidified environment that is present while imaging. This approach is feasible but is time-consuming and the atmosphere must be 100% humidified to prevent ASL evaporation. Another approach is to place oil over the culture to block ASL evaporation, but oil is not easily removed and is not O2 permeant (2). However, perfluorocarbon (PFC) is O2 permeant and evaporates with time and so it can be added to the mucosal surface during all ASL height and mucosal transport rate experiments to prevent the evaporation of ASL. PFC has been shown to have no affect on ASL height, the transepithelial voltage, or rotational mucus transport measurements and thus works well to prevent ASL evaporation (2, 7). In addition, PFC can be used as a vehicle for drug/compound additions and is used since it is immiscible with ASL (10). Given that addition of compounds in a liquid vehicle would directly increase the ASL volume, compounds are added as dry powder in a PFC suspension. By taking advantage of the fact that FC-72 PFC has a relatively low boiling point (56◦ C) compared to FC-77 PFC (97◦ C) the choice of PFC used can be tailored to the experiment. For example, we typically apply the shorter lasting FC-72 in order to measure baseline ASL volumes and for compound addition. Subsequently, FC-77 is added after addition of the compound/drug to prevent ASL evaporation.

1.4. XZ Confocal Microscopy and Measurement of the ASL

We prefer to use inverted confocal microscopes scanning in vertical (XZ) mode with high numerical aperture (1.2–1.3 NA) water or glycerol immersion lenses (see Note 6), to image from below the cultures (2). This system can obtain both qualitative and quantitative ASL data. Spinning disc confocals can also be used, although since they can only scan in the XY or horizontal mode, multiple images must be combined to make an XY stack

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and they only yield quantitative ASL data (6). We have also had some success at imaging ASL using an upright microscope (Fig. 5.3). However, high NA lenses cannot be used and instead, one must use a dry or a dipping lens (see Note 6). Furthermore, with upright microscopes, one is limited to the larger sized cultures (i.e., 30 mm diameter) since the lens cannot get close enough to the surface of the standard 10–12 mm diameter cultures. For scanning cultures on inverted microscopes, cultures are placed on a coverslip mounted in a chamber, which can either be heated or not. Several commercial designs are available (WPI and Invitrogen/Molecular Probes, etc.). The possibility also exists of using an oil lens and placing the culture directly onto the oil lens without a coverslip, but this method has not directly been tested. 1.5. Measurement of Mucus Transport Rates

The well-differentiated (ciliated) airway epithelial cultures coordinate ciliary beating to spontaneously transport mucus across the mucosal surface. This technique has revealed that the entire mucus layer moves as a single network as evidenced by the increase in linear velocity vs. the distance from the apparent center of rotation (Fig. 5.4b (2)). To measure rates of rotational mucus transport, we add 1–2 μm diameter fluorescent microspheres to the mucosal surface of the airway cultures, which preferentially segregate into the mucus layer. The microspheres are imaged in real time using conventional inverted epifluorescence microscope with a low-power dry lens and a digital camera. The mucus transport rate is then determined from 5-s exposure images (Fig. 5.4a). The movement of a single microsphere produces a streak of fluorescence in the image, with the length of the streak corresponding to the distance the microsphere has been transported in 5 s. The rotational mucus transport rates over the surface of the cultures are measured from these images, and the linear velocity of bead transport is normalized to a 1 mm distance from the center of rotation by linear regression analysis (Fig. 5.2b). Stationary mucus does not appear as streaks but can be imaged as plaques of mucus on the apical surface of the cultures (Fig. 5.4c). Alternatively, the rates of mucociliary clearance (MCC) can be determined from measurements of microsphere location changes from sequential images utilizing a high-speed CCD camera. The later technique is useful for constructing videos of the MCC (see (26)).

2. Materials 2.1. Measurement of ASL Height with Confocal Microscopy

1. Well-differentiated bronchial epithelial cell cultures. 2. Inverted confocal microscope which can perform vertical (XZ) scanning in two wavelengths simultaneously.


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Fig. 5.4. Measurement of rotational mucus transport. Epifluorescent microscopy of rotational mucus transport by airway epithelial cultures. (a) Five-second exposure image of mucus-associated fluorescent microsphere movements in a normal airway culture. (b) Plot of microsphere velocity against distance from the center of rotation (e.g., 1 mm). The slope of a best-fit line using linear regression analysis may be used to normalize transport to a set distance from the center. (c) Five-second exposure image of a stationary mucus plaque on a CF airway culture after all excess apical liquid has been absorbed by Na+ -led hyperabsorption.

Alternatively, an upright or spinning disc microscope can also be used. 3. Phosphate-buffered saline (PBS) (in mM): 125 NaCl, 4.2 KCl, 1.2 MgCl2 ·6H2 O, 1.2 CaCl2 ·H2 O, 9 K3 PO4 , pH 7.4. 4. Dextran-linked fluorescent probe. 5. Perfluorocarbon (PFC): FC-72, FC-77, or other electronic grade (3 M Company, Minneapolis, MN, USA).

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6. Ringer solution to add to the serosal side of the culture (in mM): 116 NaCl, 10 NaHCO3 , 5.1 KCl, 1.2 CaCl2 , 1.2 MgCl2 , 20 HEPES, 10 glucose, pH 7.4. 7. Cell chamber for viewing cultures on the microscope. 8. Analysis software: a. Image analysis software: e.g., ImageJ, which is freely available at http://rsb.info.nih.gov/ij/ or Metamorph b. Spreadsheet program: e.g., Excel, Microsoft, USA. 2.2. Measurement of Mucus Transport with Epifluorescent Microscopy

1. Well-differentiated (ciliated) bronchial epithelial cell cultures. 2. Inverted epifluorescent microscope with low-power lens. 3. Digital camera (either color or black and white) and image acquisition software. 4. Fluorescent microspheres (1–2 μm). 5. Perfluorocarbon (e.g., FC-77; but many other types are available with differing densities and boiling points). 6. Image analysis software (e.g., ImageJ or Metamorph).

3. Methods 3.1. Measurement of ASL (PCL and Mucus Layer) Height with Confocal Microscopy

1. Prewash cultures three times with PBS. Cultures that produce grossly visible mucus hurricanes may be excluded from ASL height measurement (see Note 1 (7)). 2. Add a PBS and fluorescent dye (0.5–2 mg/mL) to the mucosal surface of the bronchial cultures (e.g., 20 μL per 12 mm diameter culture). Then, either aspirate the excess dye to minimal ASL volumes or leave a predetermined amount of PBS/dye on the airway surface (see Notes 2 and 3). 3. Pending a suitable incubation time post-dye addition, PFC (∼100 μL) is added to the surface to prevent evaporation of the ASL and the culture is placed in the cell chamber with a serosal reservoir containing approximately 100 μL of modified HEPES-buffered Ringer solution and then placed on the scanning stage of the microscope (see Note 4). This amount of PFC was selected since it was sufficient to cover the ASL during recordings, yet evaporates soon after the cultures are returned to the highly humidified incubator (see Note 5). We have previously shown that PFC does not affect ASL height, pO2 , rotational mucus transport rates, or the transepithelial potential difference (2, 7, 9).


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4. Compounds (e.g., UTP) can be added to the ASL as dry powder suspensions in PFC. Compounds are resuspended as a slurry, either coarsely or after pulverizing in a mortar and pestle. PFC/compound slurries need to be sonicated or shaken just before addition to the cultures (see Note 5). 5. Obtain ASL heights from XZ scans of five predetermined points on each culture (one central and four circumferential) (see Note 6). 6. Import images into an image analysis software program, such as ImageJ (NIH freeware) or Metamorph (Universal Imaging). Images in a series can be opened and linked together using the stack function to speed up analysis. 7. Use the region of interest (ROI) function to measure the pixel width of the electronic magnification measurement bar from the confocal images to determine the number of pixels per micron. The scale can then be set within the program or later when analyzing in the spreadsheet. 8. Use the region of interest function to measure the height of the ASL by placing several regions of interest around the ASL that are later averaged for each image. Regions of interest are placed to avoid the 1 μm “fuzzy” interface between the ASL image and background image. Alternatively, one can perform a line scan-type analysis to measure ASL height (Fig. 5.2). 9. ASL height data acquired here are then moved to a spreadsheet, such as Excel, to analyze and determine the height. The multiple ASL heights taken for each image are averaged and then the averages for each of the five predetermined points on each culture are averaged to obtain the ASL height for that individual culture. 3.2. Fluorescent Imaging of Rotational Mucus Transport

1. Identify cultures for rotational mucus transport either visually or by phase contrast microscopy (see Note 7). 2. Add a 0.01% (vol/vol) fluorescent microsphere/PBS suspension to the mucosal surface, typically 20–50 μL per 12 mm diameter culture. 3. Add PFC to the apical surface of the cultures to prevent evaporation of the ASL. Note: The working distance of the low-power objective is adequate to image the microspheres/mucus while the cultures remain in their “longterm” culture dishes. Not removing the cultures from their dishes is advantageous as it maintains the sterility of the cultures since the dish lid of cultures does not need to be removed. 4. Using a low-power 4–10× inverted epifluorescent microscope coupled to a digital camera and data acquisition

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software, focus on the microspheres in the rotating mucus and acquire images over approximately 0.1 s, then switch to a 5-s exposure to image the “streaks” of moving microspheres. 5. Compounds suspended in PFC can then be added to the ASL. 6. Import the images into an image analysis software program. A series of images can be opened and linked together using the stack function to speed up analysis. 7. Then use the set scale function to set the size of the image based on comparison to an image of a reticule scanned under the same power. 8. Use the ROI selection tools to measure the distance (streak) the microsphere has traveled and the distance it is from the center of the hurricane. Log the data into a spreadsheet. Repeat these experiments multiple times for microspheres at varying distances from the hurricane center. 9. Plot the total distance of microsphere movement against the distance from the hurricane center, and using linear regression, extrapolate to set the distance from the center (we use 1 mm) to normalize microsphere movement. 3.3. Conclusions

In conclusion, we describe fluorescent microscopy-based methods to assess ASL structure/function. While these methods have traditionally been used to measure ASL height and mucosal transport rates, we also demonstrate that they can be employed to measure other ASL properties including the establishment of pH gradients and the molecular sieving properties of the glycocalyx. We believe these techniques will further aid in the understanding of respiratory diseases in which ASL properties are altered such as asthma, chronic pulmonary obstructive disease, and cystic fibrosis.

4. Notes 1. Cultures that spontaneously produced grossly visible mucus “hurricanes” and exhibited rotational mucus transport can be excluded from ASL height studies to remove the confounding effects of the mucus reservoir effect (7). To remove mucus from ALI cultures, approximately 0.5 mL of PBS is added mucosally and aspirated, repeating three times. In cases of excessive mucus production, the cultures may be incubated with 0.5 mL PBS at 37◦ C in the tissue culture incubator for 15 min and then aspirated. If this does not remove all mucosal mucus then the cultures can again be


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incubated with ∼0.5 mL PBS and 1 mM dithiothreitol for ∼15 min, followed by the 3× PBS rinse. 2. The fluorescent intensity of Texas Red-dextran slowly diminishes when placed in the ASL over days, due to either degradation, uptake by pinocytosis, or diffusion through the tight junctions. Hence, a greater amount of dye (2 mg/mL) will need to be added for longer duration experiments (i.e., over several days) than if they are to be examined immediately (0.2 mg/mL). 3. Depending on the experimental design, pre-washing the culture may be desirable or may obscure the response that is being measured. For example, pre-washing the cultures will remove all inhibitors of ENaC and result in a primarily absorptive phenotype (12, 14). Since there is no chemical gradient for Cl– secretion, and instead, Cl– must utilize an electrical gradient that is supplied by the inhibition of ENaC, this could be detrimental if a secretagogue was being studied under supposed physiological conditions (11). To avoid this problem amiloride (10–100 μM) or aprotinin (∼1 U/mL) can be added into the Texas Red bolus and left on the cultures. However, the natural inhibitors of ENaC that are present in the ASL are replenished within ∼24 h, so the cultures can also be washed, loaded with dye, and then used 24 h later making the addition of amiloride or aprotinin unnecessary (12, 14). 4. Our cultures are maintained under sterile conditions. However, the cultures risk exposure to pathogens (i.e., mold spores, bacteria, fungus) during all our experiments, which can destroy the cultures. The best approach to protect against infection has been to transfer the cultures from their “long-term” culture dishes (where they are kept in bronchial epithelial growth media, or BEGM) into a secondary dish containing Ringer’s solution (with PFC kept on the mucosal surface to prevent ASL evaporation). From this second dish the cultures can be transferred onto the stage of the confocal microscope. After the experiment, the cultures are washed several times in PBS and then transferred back to the “longterm” culture dish. Serial experiments over several days are thus possible but are more risky since the mucosal surface cannot be washed during the experiment since this will alter the ASL. However, with careful rinsing of the serosal surface and transference between “long-term” and temporary culture dishes after every time point we have routinely studied cultures over 5–7 days with no sign of contamination. In addition to careful handling of the cultures to minimize contamination, we also carefully wipe down our workspace around the microscope (e.g., the stage and microscope controls) and also clean the cell chamber with 70% ethanol (but

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not the lens which should only be cleaned with approved lens cleaner). When measuring ASL on multiple cultures it is recommended that the cell chamber be cleaned between either individual or small groups of cultures to prevent possible contamination from spreading. 5. Typically, the faster evaporating FC-72 will initially be added to cover the ASL, and once basal measurements have been made and the FC-72 has evaporated to minimal levels (which can be assessed visually), compounds are added to the apical surface in more FC-72. If the compounds have been added coarsely (e.g., from the packet or following grinding with a mortar and pestle), they will need to be sonicated/shaken immediately before adding them to the ASL to ensure even distribution in the PFC. An example of masses/volumes added would be to prepare 0.1 mg in 200 μL FC-72 and to aliquot a smaller amount into the ASL, such as 0.001 mg in 2 μL FC-72. 6. To image ASL by confocal microscopy, we have had the best results with a 63× water (1.2 NA) or a 63× glycerol immersion lens (1.3 NA) which has sufficient working distances (>200 μm) to image the cultures/ASL over a serosal bath with a volume 4 l), two changes per day during 3 days. 7. Sample is lyophilized.

3.1.2. Mucin Solubilization Using Protein Gel Sample-Loading Buffer

1. The sample is collected and stored with protease inhibitor (25×) at −80◦ C. 2. Reduction with DTT (final concentration of 20 mM) in protein gel sample-loading buffer at 95◦ C under magnetic stirring for 30 min, followed by 2 h at 37◦ C. 3. Alkylation by addition of IAA to a final concentration of 50 mM under magnetic stirring for 1–3 h or overnight at room temperature in the dark.

3.2. Slot/Dot Blot Followed by Visualization/ Quantification of Mucins or Large Glycoproteins 3.2.1. Slot/Dot Blotting

1. The nitrocellulose membrane is rinsed with mpH2 O and placed inside the slot/blot filtration manifold. 2. The manifold is assembled and connected to a water aspirator. 3. Samples (40–450 μl) are loaded and allowed to vacuum pull the liquid until wells are dry. 4. The membrane is developed according to Sections 3.2.2, 3.2.3, 3.2.4, or 3.2.5.

Identification and Quantification of Mucin Expression

3.2.2. Alcian Blue Staining of Mucins and Glycoproteins on Membranes (See Note 1)

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1. Wash membrane in mpH2 O for 1 min. 2. Incubate membrane in Alcian blue solution for 10–20 min. 3. Wash membrane in mpH2 O three times for 10 min. 4. Let the membrane dry. 5. Image the membrane (see Section 3.3.2).

3.2.3. PAS Staining of Mucins and Glycoproteins on Membranes (See Note 2)

1. Place the nitrocellulose membrane with the dried glycoproteins in a plastic basin. 2. Add 150 ml of mpH2 O and agitate the membrane gently at room temperature for 2 min. 3. Agitate the membrane in 150 ml of Solution A for 30 min. 4. Agitate the membrane in 150 ml of Solution B two times for 2 min. 5. Agitate the membrane in 100 ml of Schiff’s reagent for 15 min. 6. Remove the Schiff’s reagent, dry the basin quickly, and add 150 ml of Solution B and shake the membrane for 2 min. 7. Rinse membrane with mpH2 O. 8. Let the membrane dry. 9. Image the membrane (see Section 3.3.2).

3.2.4. Detection of Glycosylated Proteins with DIG Glycan Detection Kit (See Note 2)

1. The membrane is developed according to the protocol supplied with the kit.

3.2.5. Identification and Semiquantification of Mucins on Nitrocellulose and PVDF Membranes by Immunohistochemistry

1. Activate the PVDF membrane with methanol.

2. The membrane is imaged (see Section 3.3.2).

2. Incubate in blocking buffer for 1 h or overnight at 4◦ C. 3. Incubate with the primary antibody against a specific mucin (Table 8.1) at 4◦ C overnight. 4. Wash three times for 20 min in wash buffer. 5. Incubate for 2 h at room temperature with the secondary antibody. 6. Wash five times for 15 min in wash buffer. 7. Take away the major amount of solution from membrane, but avoid getting dry patches. 8. Develop the membranes with ECL according to the manufacturer’s instructions and image.

3.3. Composite Agarose– Polyacrylamide Gel Electrophoresis

The following protocol for analysis of mucins with composite agarose–polyacrylamide gel electrophoresis was originally developed by Dr. Niclas Karlsson and coworkers (4). Composite gel

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Fig. 8.2. Reduced and alkylated mucins from saliva, analyzed by composite polyacrylamide gel electrophoresis and visualized with Alcian blue. The mucins were identified by proteomics. Lane 1, whole saliva; Lane 2, saliva from submandibular/sublingual glands.

electrophoresis of mucins functions as a purification step where smaller proteins (approximately 10–100 kDa) migrate in the front. Figure 8.2 shows the analysis of reduced and alkylated salivary mucins, where MUC5B with a protein core of approximately 500 kDa separates readily from MUC7 with a protein core of approximately 100 kDa. Mucins like MUC5AC and MUC5B do not usually separate on the gel, since they are approximately of the same size. However, glycoforms can separate as we have shown for MUC5B in saliva (5), with the more acidic forms migrating further into the gel. 1. The gel casting equipment (glass plates and spacers) is mounted and placed in a 60◦ C oven, together with a gradient mixer and a pump. The pump is connected to the gradient mixer and a tubing connected to the pump is placed in-between the glass plates of the casting equipment. 2. A gel containing agarose (0.5–1% gradient), acrylamide (0–6%), and glycerol (0–10%) is prepared. The lower and upper gel solutions are prepared as described in Section 2.4, steps 10 and 11. Both solutions are boiled in a microwave oven until the agarose has melted and then immediately placed in the 60◦ C oven. 3. When the lower gel solution has cooled down to ∼60◦ C, acrylamide (40%, 1.2 ml) is added. 4. The upper and lower gel solutions (5 ml of each) are added to each chamber of the gradient mixer. APS (2 μl) and TEMED (2 μl) are added to the two chambers, respectively, and the solutions are mixed. The lower solution is stirred with a magnet and a magnetic stirrer.


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5. The pump is started at a maximum rate (10 on a GE Healthcare P-1 pump) and the two valves of the gradient mixer are opened. The gel must be casted within 2 min. 6. Two combs (2 mm × 0.75 mm) are put into the gel which is polymerized at RT for at least 6–7 h, or after 1–2 h at RT followed by +4◦ C overnight. The gels can be stored (with combs) at +4◦ C wrapped in plastic with wet papers for at least a week. 7. If the gel around the combs has shrunk, the gel can be reconstituted with the upper gel solution that has been reheated in the microwave oven. 8. Prior to electrophoresis, the combs are removed carefully, the gel is placed in the electrophoresis equipment, and running buffer is added. The wells are carefully washed with running buffer prior to use. The outer and inner containers are filled with running buffer to keep the gel cool during electrophoresis. 9. The electrophoresis container is placed on ice in a cold room (+4◦ C). After samples are loaded (sample volume limit is the well size, approximately 50–60 μl), the gel is run for 3 h at 30 mA/gel. 10. The gel is developed with a stain (see Section 3.3.1) and can then be stored in a sealed plastic bag in 1% acetic acid for at least 2 weeks. 3.3.1. Visualization of Mucins on Composite Gels

We have included three examples of stains that are all nondestructive, allowing further analysis by proteomics. Due to the dense glycosylation of mucins, these are often easily visualized with Alcian blue as there are often acidic groups present. However, for quantitative purposes, protein stains are more attractive as the type of glycosylation can vary between samples. We use Imperial stain, which is an improved Coomassie-based stain and compatible with proteomics. The commercial product SYPRO Ruby stain is a more sensitive fluorescent protein stain (in the same range as silver staining) and as this is developed for quantification of proteins in gels, this is the best for quantitative purposes. In contrast to the silver stain, this is fully compatible with subsequent proteomics.

3.3.1.1. SYPRO Ruby Staining

1. Remove the gel from the glass plates and place it in a plastic container. 2. Cover the gel with 50 ml SYPRO Ruby protein gel stain and gently agitate for 3 h or overnight. 3. Rinse gel in 10% MeOH/7% HAc for 30–60 min. 4. Wash gel in mpH2 O.

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3.3.1.2. Imperial (Coomassie) Stain

1. Remove the gel from the glass plates and place it in a plastic container and fix with 50% methanol, 10% HAc 1 h or over night. 2. Add Imperial stain and agitate the gel for 1–2 h on an orbital shaker. 3. Remove the stain and wash with mpH2 O. Destain the gel with 200 ml mpH2 O while shaking on an orbital shaker for 1–24 h.

3.3.1.3. Alcian Blue Stain

1. Fix the mucins in the gel by washing with 50% methanol and 1% acetic acid in mpH2 O for at least 1 h. 2. Equilibrate the gel by washing with 25% ethanol and 10% acetic acid in mpH2 O two times for 15 min. 3. Stain the gel with Alcian blue for 20 min. 4. Destain the gel with 50% ethanol and 10% acetic acid in mpH2 O three times for 10 min, or until satisfied. 5. Rehydrate the gel with 1% acetic acid in mpH2 O for 30 min.

3.3.2. Quantification of Mucins on Composite Gels or Blots

1. Image the SYPRO Ruby-stained gel by UV light using a fluorescent imager that allows quantification. Image the Coomassie-, PAS-, DIG-, or Alcian blue-stained gel by a visual scanner. 2. Analyze the bands by the Image J software or software coming with the fluorescent imager. Subtract background and relate the intensities to appropriate controls.

3.4. Mucin/ Glycoprotein Transfer to PVDF Membrane (See Note 3)

1. The gel is wet blotted in a Mini Trans-Blot electrophoretic transfer cell. Immobilon (PVDF PSQ ) membrane and filter papers (2×2 mm thickness) are cut out in the size of 6 cm × 9 cm. The wells of the gel are cut off before mounting the wet blot sandwich (mounting according to the manual). 2. The sandwich is placed in the transfer cell (with the gel facing the cathode), which is filled with transfer buffer and put on stirring on ice in the cold room (+4◦ C). 3. The blotting is performed at 40 W (gives ∼350–450 mA over time) for 1.5–3 h. 4. The membrane is developed with immunohistochemistry according to Section 3.2.5.

3.5. Sample Preparation for Proteomics on Mucins (See Note 4)

1. Wash the gel with mpH2 O. 2. Cut out the bands of interest on a glass plate using a scalpel. 3. Divide the gel pieces in 1.5 mm3 pieces (to facilitate enzymatic digestion) and place them into siliconized Eppendorf tubes.


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4. Destain/wash the gel plug with 0.5 ml destain solution 1 with gentle agitation for 20 min (see Note 5). 5. Destain/wash the gel plug with destain solution 2 with gentle agitation three times for 30 min or until the majority of the stain has been washed away. 6. Remove the liquid and dry the gel plugs in the speedivac. 7. Add 5–10 μl of trypsin solution to the gel pieces. If the gel plugs are not sufficiently hydrated, small aliquots of 25 mM NH4 HCO3 can be added. 8. Seal the tubes and incubate at 37◦ C overnight. 9. Add 15 μl of extract solution 1 and shake the tubes for 30 min. 10. Spin down the liquid in micro-centrifuge for a few seconds and transfer the extracted peptides to a new tube. 12. Add 20 μl of extract solution 2. 13. Shake for 30 min and pool peptide extracts. 14. Dry the peptides with vacuum centrifugation and immediately re-dissolve in 10–20 μl 0.1% HAc or HCOOH. Samples can be stored at −20◦ C. 3.6. Identification of Mucins with Proteomics

We have been more successful with nano-LC/MS than MALDI MS of tryptic peptide digests of mucins to obtain good peptide coverage. One explanation for this is that if the preparation contains the large trypsin-resistant domains, these can interfere with the crystallization on the plate. Other problems are contaminating proteins from cell debris as these can interfere with the ionization and detection of the mucin-derived peptides. It should be taken into account that most mucins generate few nonmodified tryptic peptides due to lack of frequent arginine and lysine residues and especially that many peptides do not have the expected mass due to glycan modifications. For a high-quality identification of the few non-modified peptides usually generated from mucins, each peptide should be characterized both by sequencing and with high accuracy as can be obtained by ion cyclotron resonance (ICR) or Orbitrap mass spectrometers. 1. The peptides (2 μl) are trapped on a precolumn and analyzed on the analytical column containing C18 particles. Nano-LC/MS was performed on a linear ion trap, LTQ (Thermo Finnigan). The fused silica emitter tip was held at 1.4 kV, the heated capillary at 220◦ C, and capillary voltage at 44 V. Normal scan range m/z 300–2000 (two microscans, 100 ms), data-dependent MS2 scans of the five most abundant ion in each scan, also with two microscans (100 ms), normalized collision energy of 35%, isolation window of 4.0 u, and an activation time of 30 ms.

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2. The data files obtained from nano-LC/MS of the tryptic protein digests are searched against a mucin database containing the mucin sequences that have been assembled from human and mouse genome data (see Note 6). Searches were performed using the search program MASCOT (Matrix Science).

4. Notes 1. Alcian blue is a cationic dye that binds to negatively charged carboxyl and sulfate groups on oligosaccharides. When comparing samples from different sources, it should thus be taken into account that there can be a natural variation in the amount of acidic residues, such as, for example, on salivary MUC5B in healthy individuals (5). 2. The chemistry behind both the PAS stain and the DIG glycan detection kit involves mild oxidation of vicinal hydroxyl groups in saccharide residues to aldehydes, which then are further conjugated to form a bright red dye (PAS) or to a spacer-linked steroid hapten digoxigenin (DIG). DIG is detected using an antibody conjugated to alkaline phosphatase. 3. Mucins can also be transferred to PVDF membrane using semidry blot, but large mucins with protein cores of 500 kDa or more are more efficiently transferred using wet blot. 4. Stains compatible with proteomics are the Alcian blue, Imperial stain, and SYPRO Ruby stains. The gel can be stored in 1% HAc for at least 1–2 weeks in an enclosed plastic bag, before being further processed for proteomics. Work in a clean area and work with gloves to avoid keratin contamination. 5. Alcian blue will never be removed from the gel piece. 6. The annotation and sequences of mucins in public databases have been poor, but recently these have been improved by taking advantage of our mucin database found at www. medkem.gu.se/mucinbiology/. This database contains individually assembled mucin sequences as curated by us. The database also contains assembled sequences of mucins in the FASTA format ready to be used in proteomics search engines such as MASCOT.


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Acknowledgments This work was supported by the Swedish Research Council (no. 7461, 21027, and 342-2004-4434), The Swedish Cancer Foundation, The Knut and Alice Wallenberg Foundation (KAW2007.0118), IngaBritt and Arne Lundberg Foundation, Sahlgren’s University Hospital (LUA-ALF), EU-FP7 IBDase, Wilhelm and Martina Lundgren’s Foundation, Söderbergs Stiftelser, Swedish CF Foundation, and The Swedish Foundation for Strategic Research – Innate Immunity, and The Mucosal Immunobiology and Vaccine Center (MIVAC). References 1. Garcia, M. A., Yang, N., and Quinton, P. M. (2009) Normal mouse intestinal mucus release requires cystic fibrosis transmembrane regulator dependent bicarbonate secretion. J Clin Invest 119, 2613–2622. 2. Caramori, G., DiGregorio, C., Carlstedt, I., Casolari, P., Guzzinati, I., Adcock, I. M., et al. (2004) Mucin expression in peripheral airways of patients with chronic obstructive pulmonary disease. Histopathology 45, 477–484. 3. Larsson, J. M., Karlsson, H., Sjövall, H., and Hansson, G. C. (2009) A complex, but uniform O-glycosylation of the human MUC2 mucin from colonic biopsies analyzed by nanoLC/MSn . Glycobiology 19, 756–766. 4. Schulz, B. L., Packer, N. H., and Karlsson, N. G. (2002) Small-scale analysis of O-linked oligosaccharides from glycoproteins and mucins separated by gel electrophoresis. Anal Chem 74, 6088–6097. 5. Thomsson, K. A., Schulz, B. J., Packer, N., and Karlsson, N. G. (2005) MUC5B glycosylation in human saliva reflects blood group and secretor status. Glycobiology 15, 791–804. 6. Schroeder, J. A., Thompson, M. C., Gardner, M., and Gendler, S. J. (2001) Transgenic

7.

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MUC1 interacts with epidermal growth factor receptor and correlates with mitogenactivated protein kinase activation in the mouse mammary gland. J Biol Chem 276, 13057–13064. Godl, K., Johansson, M. E. V., Karlsson, H., Morgelin, M., Lidell, M. E., Olson, F. J., et al. (2002) The Ntermini of the MUC2 mucin form trimers that are held together within a trypsinresistant core fragment. J Biol Chem 277, 47248–47256. Lidell, M. E., Bara, J., and Hansson, G. C. (2008) Mapping of the 45M1 epitope to the C-terminal cysteine-rich part of the human MUC5AC mucin. FEBS J 275, 481–489. Thornton, D. J., Gray, T., Nettesheim, P., Howard, M., Koo, J. S., and Sheehan, J. K. (2000) Characterization of mucins from cultured normal human tracheobronchial epithelial cells. Am J Physiol 278, L1118–L1128. Bast, R. C., Feeney, M., Lazarus, H., Nadler, L. M., Colvin, R. B., and Knapp, R. C. (1981) Reactivity of a monoclonal antibody with human ovarian carcinoma. J Clin Invest 68, 1331–1337.

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Chapter 9 Methods to Classify Bacterial Pathogens in Cystic Fibrosis Thomas Bjarnsholt, Xiaohui Chen Nielsen, Ulla Johansen, Lena Nørgaard, and Niels Høiby Abstract Many bacteria can be detected in CF sputum, pathogenic and commensal. Modified Koch’s criteria for identification of established and emerging CF pathogens are therefore described. Methods are described to isolate bacteria and to detect bacterial biofilms in sputum or lung tissue from CF patients by means of conventional culturing and staining techniques and by the PNA FISH technique. Additionally, the confocal scanning laser microscopy technique is described for studying biofilms in vitro in a flow cell system. The recA-gene PCR and the RFLP-based identification methods are described for identification of isolates from the Burkholderia complex to the species level. DNA typing by PFGE, which can be used for any bacterial pathogen, is described as it is employed for Pseudomonas aeruginosa. A commercially available ELISA method is described for measuring IgG antibodies against P. aeruginosa in CF patients. Key words: Biofilm, Koch’s criteria, Pseudomonas aeruginosa, Burkholderia, FISH, Pseudomonas antibodies, pulsed field gel electrophoresis, confocal scanning laser microscopy, recA-gene PCR, microbiology.

1. Introduction The lower respiratory tract of normal persons and CF patients does not harbor a normal microbiological flora. On the other hand, aspiration of secretion from the pharynx occurs during, e.g., sleep, and exposure to aerosols also leads to repeated daily exposure of the defense system of the lungs to microbes carried by droplets, droplet nuclei, and minor solid particles (1). Due to the basic defect in CF which implies reduced volume of the periciliary fluid of the respiratory mucosa, the non-inflammatory mucociliary clearance of inhaled microorganisms is impaired, and the patients therefore have to recruit the inflammatory defense M.D. Amaral, K. Kunzelmann (eds.), Cystic Fibrosis, Methods in Molecular Biology 742, DOI 10.1007/978-1-61779-120-8_9, © Springer Science+Business Media, LLC 2011

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mechanisms which may imply inflammation (2, 3). When respiratory secretion is obtained for routine detection of supposed CF pathogens, such secretions are nearly always contaminated by microbes from the pharynx and mouth, where aerobic, facultative, and anaerobic bacteria are part of the normal flora (4, 5). Actually, any mouth contains at least 100–200 different bacterial species (not all can be cultured) and the number of bacterial cells is approximately 108 /ml saliva. Well-established CF pathogens such as Staphylococcus aureus, Haemophilus influenzae, Streptococcus pneumoniae, and Moraxella catarrhalis may be members of the normal flora (4). Other CF pathogens such as Pseudomonas aeruginosa, Burkholderia cepacia complex, Stenotrophomonas maltophilia, Achromobacter xylosoxidans, and MOTT are not members of the normal human flora but are environmental bacteria which may occur in water (including tap water), soil, and even food (6). The fungus Aspergillus fumigatus (7) is also an environmental microorganism which is found in humid niches in buildings, e.g., bathrooms, kitchens, and air ventilation tubes. The major problem is therefore not to detect microbes in the respiratory secretions from CF patients but to identify whether the detected microbes originate from the lower respiratory tract and whether they are causing any damage (pathogens) or rather are innocent bystanders which only contaminate the secretions (sputum) from the lower respiratory tract on its way through the major bronchi. This is of clinical importance, since sputum is a waste product located outside the body’s mucosa – like stools – but it may contain pathogens which are also present in the lower respiratory tract – the respiratory zone of the lungs – and contribute to the pathogenesis of the tissue damage in the lungs of the CF patient. It is therefore important to identify the characteristic features of those microorganisms which play a role in the pathogenesis of the lung pathology of CF patients. It is also important to realize that sputum of CF patients is anaerobic and anaerobic bacteria may play a role during exacerbations of the CF lung symptoms (8–10). 1.1. What Is a CF Pathogen?

In the golden era of bacteriology 130 years ago where many of the major and specific pathogens were detected, confusion of pathogens and contaminants was common but the famous postulates of Koch-Henle (11) were very helpful in the identification of the etiology of “specific” infectious diseases like tuberculosis: (1) The organism must be present in every case of the disease in question and under circumstances which can account for the pathological changes and clinical course of the disease. (2) The organism occurs in no other disease as a fortuitous and non-pathogenic parasite. (3) After being fully isolated from the body and repeatedly grown in pure culture it can induce the disease anew.

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Obviously these postulates did not work on “non-specific” infectious diseases like pneumonia, where several microbes may give rather similar symptoms and pathology. The postulates have therefore subsequently been modified (12): (1) Epidemiology: The organism should be detected more frequently and/or in larger numbers from patients with disease than in those without; (2) Antibody: an antibody response measured by any of several available techniques, should be demonstrated in the infected host; (3) Response to treatment: Clinical and microbiological cure after treatment with an antimicrobial agent to which the organism is susceptible in vitro; (4) Transmissibility: The organism should infect an animal host from which they can be recovered and, in so doing, produce disease similar to that seen in man. Based on lessons learned from P. aeruginosa in CF, further modifications of the postulates are necessary to identify criteria which should be fulfilled before isolated microbes are considered CF pathogens (6): (1) Correct species identification of suspected pathogens – every time it is isolated from the patient. Chronic infections may not be diagnosed if the microbe’s name is changing from time to time in the same or different laboratories (e.g., Gramnegative rods → P. aeruginosa → non-fermenter). Even change of taxonomic classification may give rise to such diagnostic errors (e.g., Pseudomonas maltophilia → Xanthomonas maltophilia → S. maltophilia) in a patient who actually fulfilled the criteria of chronic infection by culture (continuous presence of the bacteria for ≥6 months based on monthly visits or >50% positive cultures during 1 year based on visits every 3 months) and/or increased antibody response (13–16). Furthermore, phenotypic variations of colony morphology are common in CF, so each phenotype should be identified and its antibiotic susceptibility should be examined (e.g., mucoid, non-mucoid, and small colony variants of P. aeruginosa; Fig. 9.1) (2). Are the identified microbes causing inflammation in the CF airways and decrease of pulmonary function and eventually lung transplantation or death, or is it an innocent bystander to another established CF pathogen which is also present, e.g., S. aureus? (3) Are there any clinical and bacteriological effects of antibiotic therapy to which the organism is susceptible in vitro or is it an innocent bystander to another established CF pathogen, e.g., S. aureus, which is also present and also susceptible to the antibiotic therapy? It is, however, important to realize that biofilm-growing bacteria such as mucoid P. aeruginosa are generally not eradicated by antibiotic therapy, although their number may decrease for a period of time. In spite of that, a clinical response to antibiotic therapy is regularly seen (4). Epidemiology: Is there any spread of the microorganism to other CF patients as determined by genotyping of the isolates? (5) Antibody response: chronic infections in CF are characterized by a pronounced and diagnostic important

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Fig. 9.1. Culture (72 h, 37◦ C, modified Conradi–Drigalski medium) selective for Gramnegative rods (blue plates) of sputum from a CF patient with chronic P. aeruginosa lung infection. Top arrow: Mucoid phenotype (alginate producing), middle arrow: non-mucoid phenotype, bottom arrow: small colony variant. All three phenotypes had the same PFGE DNA type = the same clone.

antibody response but cross-reactive antibodies are found in CF, so the specificity of the antibody response may be questioned and absorbance experiments carried out (16) (6). Although the microbes may possess recognized virulence factors and toxins, they may not play a role during chronic infection because of induction of neutralizing antibodies (17). Actually during chronic infections, especially if biofilm-growing bacteria (e.g., P. aeruginosa) are involved, immune complex-mediated inflammation dominates the pathology of the lung tissue damage (18). 1.2. Isolation and Identification of Pathogenic Bacteria from the CF Lung 1.2.1. Microscopy of Gram-Stained Specimens

Sputum can be obtained by coughing or, in non-sputum producers, by deep throat culture, endolaryngeal suction, and bronchoscopic lavage (4). Obviously, bronchoscopy is not suitable for routine screening purpose, although the risks are small (19). Results obtained by that method and deep throat culture simultaneously indicate that the predictive values of throat cultures positive for S. aureus or P. aeruginosa were 91 and 83%, respectively, whereas the negative predictive values were 80 and 70%, respectively (4). In the Danish CF Centre in Copenhagen, tra-


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cheal secretion is obtained by endolaryngeal suction and the bacteriological examination of the secretion comprises microscopy and culture of the secretion. Gram-stained films of the specimens are examined by microscopy and the bacterial flora associated with areas consisting of respiratory epithelial cells and polymorphonuclear leukocytes and mucus but without squamous epithelial cells is described, whereas the flora associated with oral epithelial cells is considered to be predominantly of oral or pharyngeal origin, and not described (4). Based upon these criteria, 85% of the culture-positive samples (P. aeruginosa 91%, S. aureus 81%, H. influenzae 87%, and S. pneumoniae 85%) were in accordance with the results obtained by microscopy (4). A presumptive bacteriological diagnosis can frequently be obtained by the microscopic examinations (Fig. 9.2) as a guideline for urgent chemotherapy if this is indicated, but in most cases the chemotherapy can be withheld until the results of the culture are obtained. Indication for antimicrobial chemotherapy includes that the bacteria in question have been cultured as well as found by microscopy according to the principles described above, since microscopy of Gram-stained smears is the best routine method to distinguish between oral contaminants and microbes present in secretions from the lower respiratory tract (4). 1.2.2. Media and Culture Conditions

The media used for culturing the bacteria from CF sputum include the following (1): enriched standard media, since auxotrophic mutants requiring specific amino acids as growth factors are common in CF (4), e.g., chocolate agar and horse or sheep blood agar; (2) selective media from Gram-negative enteric and non-fermentative bacteria (Enterobacteriaceae, Pseudomonas species, Achromobacter species, S. maltophilia); (3) selective media (mannitol salt agar) containing 7.5% NaCl for S. aureus (20); furthermore, for detection of methicillin-resistant S. aureus (MRSA), several chromogenic media are on the market such as MRSA ID (BioMerieux, France) (21). When distinct green colonies (MRSA) are grown, then the mecA gene can be detected by PCR or DNA hybridization techniques using, e.g., mecA EVIGENETM (AdvanDx, MA, USA) (22). (4) Selective media containing colistin for Burkholderia species (4, 6). (5) In some centers also selective media for H. influenzae, Sabouraud agar for fungi (e.g., A. fumigatus), and Löwenstein–Jensen medium for mycobacteria other than tuberculosis (MOTT), but decontamination of the secretion with 0.25% N-acetyl-L-cystein and 1% NaOH followed by 5% oxalic acid is necessary to detect the mycobacteria in the presence of, for example, P. aeruginosa (4, 6). In the Danish CF Centre in Copenhagen, a blood agar plate for primary sensitivity testing with antibiotics (tablets or disks) active to P. aeruginosa (colistin, tobramycin, piperacillin+tazobactam, ceftazidime, aztreonam, meropenem, and ciprofloxacin) is also

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Fig. 9.2. Gram-stained smears from three CF patients with chronic P. aeruginosa (all had mucoid phenotypes) lung infection. The red arrows show biofilms of P. aeruginosa and the black arrows show polymorphonuclear leukocytes. Notice the great diversity of the appearance of the biofilms. Magnification 1000×.

used to detect minority populations of resistant mutants in the presence of a majority of susceptible phenotypes. Incubation takes place for 72–96 h at 37◦ C (6 weeks for mycobacteria, but frequently MOTT isolated from CF patients are rapid growing (Mycobacterium chelonei, Mycobacterium fortuitum) and colonies are detected within 1 week). The chocolate agar medium is incubated in 5% CO2 atmosphere. Anaerobic culture is generally not


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useful unless quantitative cultures are used or if the secretion is obtained by bronchial alveolar lavage, since heavy growth of the normal anaerobic flora from the mouth. The isolated bacteria are then identified to the species level using standard biochemical tests and secondary sensitivity testing to relevant antibiotics is carried out. The phenotypic appearance of the colonies (e.g., mucoid and non-mucoid and small colony variants of P. aeruginosa; Fig. 9.1) is described in the report from the laboratory and the antibiotic susceptibility of each phenotypic variant is determined. It is important to realize that notably P. aeruginosa phenotypes change during the chronic lung infection, leading to changed LPS (rough or semi-rough), small colony variants, auxotrophy, loss of pigment or changed pigment production, slow growth, and changed phenotypic colony morphology (Fig. 9.1) (e.g., mucoid colonies) (4, 6, 23, 24). The identification of such phenotypic variants may therefore be difficult leading to misclassification (25, 26). Classical biochemical tests not included in the commercial identification panels may therefore often be necessary to employ for identification of the species. Newer methods based on the profile of proteins of the bacteria detected directly from an intact bacterial cell surface by matrix-assisted laser desorption ionization time-of-flight mass spectrometry (MALDI-TOF) (27) or sequencing of the 16S rDNA gene may therefore be indicated in uncertain cases (28). In order to investigate suspected epidemiologically related isolates, genotyping by, e.g., PFGE or other DNA-based typing methods is necessary (29). Chronic infection of P. aeruginosa or other pathogens is defined by the Copenhagen criteria (4) or by Leed’s criteria (15) but requires 6–12 months observation time. However, detection of a significant rise of specific anti-Pseudomonas IgG antibodies shows about 90% predictive value of a positive test and 90% predictive value of a negative test ability to detect chronic infection with these bacteria (16). 1.3. Identification of Pathogenic Bacteria from the CF Lung – Molecular Biology Methods vs. Conventional Methods

For chronic infections, it can be very problematic to isolate pathogenic bacteria. An exception is actually CF in which the easily accessible purulent sputum coughed up by the patients on a regular basis harbors the bacteria. The next problem which also applies for CF is the identification of the bacteria present. In the clinical microbiology laboratory, many techniques and procedures are available for identifying the bacteria such as microscopy, culture, PCR, and sequencing of 16S rRNA gene. Each method has its advantages and disadvantages. For culturing, the problem is to collect and grow the offending pathogen. Since sputum is coughed up through the trachea and oral cavity, oral bacteria might be abundantly present as well. Also the bacteria of the chronic CF lung infections may have very slow growth rate (30) in vivo and

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in vitro. Due to these possible contaminations, one has to analyze cultures of CF sputum very carefully. On the other hand, if bacteria are cultured, both identification and susceptibility to antibiotics can be done. The usage of molecular techniques such as PCR will detect even tiny amounts of DNA or RNA available in the sample (31). Here again contamination of the oral flora has to be avoided. The second problem is to avoid eliminating the minor fraction of prokaryotic DNA/RNA by the eukaryotic DNA/RNA, which might either interfere with the PCR reaction (fairly easy to solve) or outcompete the capacity for DNA/RNA extraction. Another problem with PCR is that every DNA/RNA fragment homologous to the primers will be detected, including defragmented DNA/RNA, which may generate false positives. Additionally, just because a bacterium is present does not necessarily indicate that it contributes to the pathogenesis of the infection (see above), so it might not need to be treated. The final problem which may be the biggest challenge in the coming years for biofilm infections in CF patients is to reveal the significance of the numerous microorganisms detected by molecular techniques (see above). 1.4. Visualization of Bacteria in Sputum and Lung Tissue

Chronic bacterial infections are caused by the ability of bacteria to organize themselves in micro-colonies, also called biofilms (3, 32–36). In this state, the bacteria are imbedded in a self-produced protective matrix, often with surrounding inflammatory cells (36, 37). Bacteria living in biofilms are very well protected against antibiotics and the host defense. Due to this, it is a necessity to consider whether the isolated and identified pathogenic bacteria were as single, planktonic cells, which are easy to treat, or as difficult-to-treat biofilms, which is the problem in CF and other chronic infections. Microscopy enables direct visualization of the infecting bacteria (Fig. 9.2), but in case of the presence of several similar looking bacteria (Gram-negative rods), identification of the bacteria in, e.g., a biofilm in sputum is not always possible. This can be done using fluorescent antibody probes (3, 8) or DNA probes for FISH (3), see later.

1.5. In Vitro Study of Biofilms

The developmental processes of biofilms have been thoroughly studied in surface-based in vitro systems and are relevant for studies on CF pathology and microbiology (3). The most studied bacterium in this context is P. aeruginosa. The ability of P. aeruginosa to form biofilms is thought to be one of its main survival strategies in an infectious process and is considered an important pathogenicity trait. The in vitro-formed biofilm consists of micro-colonies encapsulated by exopolysaccharide (EPS) produced by the bacteria


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itself, though most of the biofilms are made up of water channels which are thought to operate as a distribution system of nutrients and oxygen. An oxygen gradient is present from the surface decreasing downward to the substratum (8, 38). Pseudomonas aeruginosa forms biofilms on almost any surface and any condition, both nutritional and environmental. From time to time, it is stated that biofilm formation, from planktonic to sessile mode of growth, is a complex and highly regulated process (39). This slight variation in biofilm structure by the same strain of P. aeruginosa indicates that biofilm formation, and the successive growth and development, is a complex but somewhat arbitrary process. It has also been suggested that biofilm formation is dependent on the expression of a specific biofilm program (40). However, based on all the in vivo observations present today, it is more likely that biofilm formation proceeds through a series of temporal events that probably reflect adaptation to nutritional and environmental conditions (41–43). The in vitro, surface-based, biofilm developmental process can be divided into different stages: (i) attachment, (ii) maturation, and (iii) dispersion, as indicated by Sauer et al. (44) and Klausen et al. (45).

2. Materials 2.1. Biofilm Detection in Sputum and Tissue (3, 4, 46) 2.1.1. Gram Stain

2.1.2. PNA FISH of Tissue and Sputum

Material for investigation has to be either (a) fresh sputum or (b) section of lung tissue and the following: 1. Microscope slides. 2. Gram stains (crystal violet for staining, Lugol’s iodine as mordant, 99% ethanol for decolorizing, and basic carbol fuchsin for counterstaining) (20). 1. Material for investigation has to be either (a) fresh sputum or (b) sections of lung tissue: a. For paraffin-embedded tissue sections: i. Xylene. ii. 99.9% ethanol. iii. 96% ethanol. iv. Sterile water. 2. Microscope slides for fluorescence microscopy. 3. Cover slips, thickness 0.15 mm. 4. Staining dish with slide holder. 5. Heating block (55◦ C).

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6. Water bath (55◦ C). 7. PNA FISH probes (AdvanDx, MA, USA). 8. 60× wash solution (AdvanDx, MA, USA). 9. Vectashield mounting medium with DAPI (Vector Laboratories, Burlingame, CA, USA). 10. Fluorescence microscope equipped with a 60–100× oil objective, camera for fluorescence, and the correct filters for the probes to be used. 11. Immersion oil. 2.2. In Vitro Flow Cell Biofilm

1. Bacterial strains: The investigated bacteria have to express a fluorescent protein (like GFP) by genetic manipulation or staining with a DNA stain. 2. AB trace minimal growth medium: Add 1 mM MgCl2 , 0.1 mM CaCl2 , and 100 μL/L trace metals (see description later) to water and sterilize the solution by autoclaving. Thereafter add 10% A10 (see description later). Media should be heated to 37◦ C before use. 3. Trace metal solution: 200 mg/L CaSO4 ·2H2 O, 200 mg/L FeSO4 ·7H2 O, MnSO4 ·H2 O, 20 mg/L CuSO4 ·5H2 O, 20 mg/L ZnSO4 ·7H2 O, 10 mg/L CoSO4 ·7H2 O, 12 mg/L Na2 MoO4 ·H2 O, and 5 mg/L H3 BO3 . Can be stored at room temperature for several months. Remember to mix before use. 4. A10: Add 20 g (NH4 )2 SO4 , 60 g Na2 HPO4 , 30 g KH2 PO4 , and 30 g NaCl to 1 L water. Adjust pH to 6.4 and sterilize solution by autoclaving. Store at 4◦ C. 5. Glucose. Use 0.3 mM. 6. BactoTM casamino acid. 7. 16-Channel Watson Marlow 205S peristaltic pump. 8. Equipment to assemble the system: 8.1. A: Silicon tube, internal diameter: 2 mm, external diameter: 4 mm (Ole Dich, Denmark). 8.2. B: Silicon tube, internal diameter: 1 mm, external diameter: 3 mm (Ole Dich, Denmark). 8.3. 1: Straight connectors 1/8 in. (Buch & Holm, Denmark). 8.4. 2: T-connectors 1/8 in. (Buch & Holm, Denmark). 8.5. 3: Reducing connectors 1/8 in. × 1/16 in. (Buch & Holm, Denmark). 8.6. 4: Barrel tip cap orange (Diatom, 5113-XM). 9. Silicone to assemble the flow chamber to the cover slip.


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10. Surface-attached mono species biofilms are cultivated in flow chambers with channel dimensions of 1 mm × 4 mm × 40 mm. 11. The substratum consists of a microscope cover slip (24 mm × 50 mm). 12. For sterilization of biofilm system: Use sodium hypochlorite (NaClO) solution (13%), bleach, and dilute it in water to 0.5% (Note 1). 2.2.1. Confocal Scanning Laser Microscopy

2.3. recA-Gene PCR and RFLP-Based Identification Approach for B. cepacia Complex

Image acquisitions of biofilms are performed using a confocal scanning laser microscope (CSLM) such as Leica SP5 (Leica Lasertechnik, GmbH, Heidelberg, Germany) equipped with a detector and a filter set for monitoring GFP and propidium iodide (PI). In addition, a reflection detector for acquiring bright-field images is installed. Images should be obtained using a 40–100× oil objective. Image scanning is carried out with the 488-nm laser line from an Ar/Kr laser. Imaris software package (Bitplane, AG) can be used to generate pictures of the biofilm. 1. MagNa Pure LC DNA Isolation Kit III (Bacteria, Fungi) (Roche). 2. dNTP (10 mM; Qiagen Laboratories, Inc.). 3. PCR buffer, 10× (Qiagen Laboratories, Inc.). 4. Q-solution, 5× (Qiagen Laboratories, Inc.). 5. Taq polymerase (Qiagen Laboratories, Inc.). 6. Sterile distilled water. 7. Primers for B. cepacia complex-specific PCR: BCR 1: 5 -TGA CCG CCG AGA AGA GCA A-3 (50 μM; DNA Technology, Aarhus, Denmark). BCR 2: 5 -CTC TTC TTC GTC CAT CGC CTC-3 (50 μM; DNA Technology, Aarhus, Denmark). 8. Primers for Burkholderia multivorans-specific PCR: BCRBM 1: 5 -CGG CGT CAA CGT GCG GGA T-3 (50 μM; DNA Technology, Aarhus, Denmark). BCRBM 2: 5 -TCC ATC GCC TCG GCT TCG T-3 (50 μM; DNA Technology, Aarhus, Denmark). 9. PCR mastermix (per sample): 1.25 μl dNTP, 2.0 μl primer 1, 2.0 μl primer 2, 5.0 μl 10× PCR buffer, 10.0 μl 5× Q-solution, 0.5 μl Taq polymerase, 19.25 μl sterile distilled water. 10. Agarose (Sigma A-4010 low gelling). 11. HaeIII restriction enzyme (10 units/μl; New England Biolabs, Inc., MA, USA).

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12. TBE buffer 10× conc. (27 mM Tris base, 27 mM boric acid, 0.75 mM EDTA, pH 8.5). 13. Ethidium bromide (AMRESCO, Inc., Cleveland, OH, USA). 14. Loading buffer: TE EC-1028 (orange sample buffer; EmpiTec, CA, USA). 15. Molecular marker (DNA 100-bp ladder, N3231S) (New England Biolabs, Inc., MA, USA). 16. Molecular marker (DNA 50-bp ladder N3236S) (New England Biolabs, Inc., MA, USA). 17. Criterion TBE ready polyacrylamide gel 10% (Bio-Rad, CA, USA). 2.3.1. Instruments

1. MagNa Pure LC (Roche, Switzerland). 2. FlexCycler (AH Diagnostics). 3. Bio-Rad PowerPac Basic and Criterion Cell. 4. Agarose electrophoresis apparatus Power Pack Base (BioRad, CA, USA). 5. Gel Doc (Bio-Rad, CA, USA). 6. Galaxy minister centrifuge (VWR Instruments, Korea).

2.4. DNA Typing by Pulsed Field Gel Electrophoresis (PFGE) for Typing of P. aeruginosa or Other Species (29, 50)

1. SE buffer (75 mM NaCl, 25 mM EDTA, pH 7.4). 2. Agarose (Sigma A-4010 low gelling). R Gold agarose (BioWhittaker Molecular Applica3. SeaKem tions, Inc., Rockland, ME, USA).

4. ES buffer (1% (w/v) N-lauroylsarcosine, 0.5 M EDTA, pH 9.5). 5. Proteinase K (AMRESCO, Inc., Solon, OH, USA). 6. Spe1 restriction enzyme (New England Biolabs, Inc., MA, USA). 7. Bovine serum albumin (Sigma A 7030 albumin). 8. Tris–borate buffer (27 mM Tris base, 27 mM boric acid, 0.75 mM EDTA, pH 8.5). 9. TE buffer with bovine serum albumin (BSA) (50 mM potassium acetate + 20 mM Tris–acetate + 10 mM magnesium acetate + 1 mM DTT, pH 7.9 + 2.5 mg BSA/ml). 10. Thiourea (thiocarbamide, 99%; Bie & Bernsen, Denmark). 11. Ethidium bromide (AMRESCO, Inc., Cleveland, OH, USA). 12. TRE buffer (10 mM Tris base + 10 mM EDTA, pH 7.4).


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13. NEBuffer 4 (50 mM potassium acetate + 20 mM Tris– acetate + 10 mM magnesium acetate + 1 mM DTT, pH 7.9; BioLabs, MA, USA). 2.5. ELISA for Measuring IgG and IgA Antibodies Against Bacterial Sonicate (16, 53)

Pseudomonas-CF-IgG ELISA kit (Statens Serum Institute, Copenhagen, Denmark) contains the following: 1 MaxiSorp R ), 9 mg lyophilized antigen (sonication of ELISA plate (NUNC P. aeruginosa serotypes O-1 through O-17), 1 vial pooled human standard antiserum (antibodies against P. aeruginosa), 0.5 ml sterile distilled water, 12 ml coating buffer, 250 ml washing buffer, 250 ml dilution buffer, 0.1 ml rabbit–anti-human IgG HPR, 12 ml sulfuric acid (2 M), 12 ml TMP plus standard, and ELISA reader set at 450 nm.

3. Methods 3.1. Gram Stain 3.1.1. Sputum

1. A thin smear is produced by gently spreading purulent sputum on top of the microscope slide by a loop and is left for air-drying. 2. The slide with the dried smear is then gently heat fixated. 3. Gram staining is performed according to the routine of the laboratory. 4. Using immersion oil and 1000× magnification, aggregated bacterial cells which appear to be surrounded by a self-produced matrix are searched. There is a great variation in the size and shape of biofilms and the specimen has to be examined for 5–10 min in order to detect possible biofilms. Mucoid P. aeruginosa regularly produce detectable biofilms in sputum which resemble smears of mucoid colonies from solid media (23). Biofilms of A. xylosoxidans, B. cepacia complex, and S. maltophilia can also be detected, but such biofilms appear more condensed compared to biofilms of mucoid P. aeruginosa (46). Non-mucoid P. aeruginosa do not appear to produce biofilm in vivo in CF (3).

3.2. PNA FISH of Tissue and Sputum 3.2.1. Sputum 3.2.1.1. Tissue Sections

1. A thin and uniform smear, of the freshly isolated sputum, is made on a microscope slide (intended for fluorescence). 2. The material is heat fixated. Before hybridization of paraffin-embedded tissue, paraffin has to be removed from the tissue sections.

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De-paraffinization protocol 1. Immerse slides in xylene (2 × 5 min). 2. Immerse slides in 99.9% ethanol (2 × 3 min). 3. Immerse slides in 96% ethanol (2 × 3 min). 4. Immerse slides in water (3 × 3 min). 5. Air-dry. Slides are stable for months at RT. 3.2.2. PNA FISH

The de-paraffinized tissue sections and the sputum smears were analyzed by fluorescence in situ hybridization (FISH) using peptide nucleic acid (PNA) probes. In a recent study, we used a mixture of Texas Red-labeled, P. aeruginosa-specific PNA probe and fluorescein-labeled, universal bacterium PNA probe (AdvanDx, MA, USA) (3). 1. The probe of choice is added to each section or sputum smear. 2. A cover slip is placed on top of the material; a pencil is used gently to spread the probe under the slide and to remove bubbles. 3. The microscope slide is placed on a heating block for hybridization at 55◦ C for 90 min covered by a lid. 4. The slides are washed for 30 min at 55◦ C in wash solution (AdvanDx). 5. Vectashield mounting media with 4 ,6-diamidino-2phenylindole (DAPI) (Vector Laboratories, Burlingame, CA) are applied, and a cover slip is added to each slide; a pencil is used gently to spread the probe under the slide and to remove bubbles. 6. Slides are inspected using a fluorescence microscope equipped with a camera and appropriate filters like a fluorescein isothiocyanate (FITC), a Texas Red, a DAPI, a dual FITC/Texas Red, and a dual DAPI/Texas Red filter (Fig. 9.3) (3) (Note 2).

3.3. In Vitro Flow Cell Biofilm

Using an in vitro continuous culture flow cell system (47) enables continuous non-invasive investigation of biofilm development. This biofilm flow cell system is perfect for visual inspection of formation, disruption, and killing of biofilms using a CSLM. The biofilm is monitored using either fluorescent-tagged bacterial cells or a DNA stain such as Syto9. The killing of the bacteria is monitored using PI which will stain the DNA of cells with impaired membrane, i.e., dead cells. PI fluoresces red which is why another color such as green is needed for the live bacteria.


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Fig. 9.3. Pseudomonas aeruginosa biofilms surrounded by PMNs visualized by PNA FISH and DAPI, located intraluminally in the conductive zone of the lung of a CF patient.

1. Assemble the flow cells for minimum 24 h prior to use by gluing (use silicone) a 24 mm × 50 mm glass cover slip onto the top of the flow cell (see Fig. 9.4B). 2. Assemble the rest of the system as shown in Fig. 9.4A. 3. The following description is based on a 16-channel pump from Watson Marlow (205S). Sterilize the whole system using 1 L of 0.5% NaClO solution in sterilized Milli-Q water. Set the pump at 90 rpm to fill the whole system. When the bobble traps are filled, put on the caps and set the pump at 12 rpm. Sterilize for approximately 2 h. Make sure that nothing is leaking, otherwise use silicone to stop it. Remember to wear gloves and glasses when working with NaClO. 4. Drain the system for liquid setting the pump at 90 rpm. 5. Wash the system two times with 1 L of sterilized water in the same way as with the NaClO. Set the pump at 5– 50 rpm when the system is filled. Empty the system after one 1 L and fill up again with the new flask (see Note 3). 6. After the last wash, drain the system and fill with AB trace minimal medium. Place the system at 37◦ C overnight (see Note 3). 7. Make an overnight culture of the bacteria to be investigated.

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Fig. 9.4. A: In vitro continuous culture biofilm flow cell system setup. The numbers and letters refer to the designation in the material section of the different connectors and tubes. B: Three-channel flow cell, in which the biofilms are cultivated. Confocal scanning laser microscopy.

8. Before inoculation with the overnight culture, stop the flow and clamp off the tubes between the flow channels and the bubble traps. 9. Inoculate the flow chambers using 250 μL of the overnight culture diluted to an OD600 of 0.001–0.1 in 0.9% NaCl. Inject the diluted culture in the flow channels by using a syringe needle which is inserted into the tubing next to the flow channel inlet. Close the injection hole with a thin layer of silicone. 10. Arrest the medium flow for 30–60 min to allow efficient bacterial attachment to the glass surface. Turn the flow cell upside down placing it on the glass surface. 11. Turn the flow cells again so that the glass slides are facing up. Remove the clamps and start the medium flow. The medium should be pumped at a constant rate of 1.75 rpm for a 16-channel pump and 2.00 rpm for a 12-channel pump which corresponds to approximately 3.3 ml/h using the peristaltic pumps from Watson Marlow (205S). 12. Let the biofilm develop. We allow the biofilm to develop and mature in the flow chambers for 3–4 days for antibiotic tolerance investigations.


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13. At the day of treatment, the growth medium is changed to fresh growth medium containing antibiotics or other relevant substance. The medium is changed by (1) stopping the flow (2); clamping of the tube between the flow cells and bubble traps (3); emptying the bubble traps by pulling the syringe off and quickly again (4); removing the barrel tip cap and filling the bubble traps with the new growth medium containing antibiotics, e.g., at 90 rpm (5); stopping the system when the bubble traps are filled up and removing the clamps; and (6) starting the system again at 1.75 rpm. 14. Investigate the biofilm approximately at any given time point relevant to the treatment. 15. For examination of the biofilm with CSLM, a viability staining kit is used: If a non-GFP-tagged P. aeruginosa strain is used, LIVE/DEAD viability staining kit can be applied (Fig. 9.5). Syto 9 (Invitrogen, cat. no. S-34854) and PI (Sigma, P-4170) are added at a concentration of 0.005 and 0.01 mM, respectively, 15 min before examination of the flow cells by injecting it in the same way as the bacterial culture explained in Section 9–10 (DNA stains are often light sensitive, which is why the system should be shielded from light – see Note 4). 16. If a GFP-tagged P. aeruginosa strain is used, only PI is added to the flow chambers. 3.4. recA-Gene PCR and RFLP-Based Identification Approach for B. cepacia Complex

The B. cepacia complex is a very diverse group of bacteria. RecA gene has proven to be useful for the identification of B. cepacia complex species (26, 48). The method described includes three steps: Step 1. PCR amplification of B. cepacia complex recA gene. By applying primers (BCR1 and BCR2) that are specific for B. cepacia complex, a single 1,043-bp amplicon will be generated from all strains representative of the B. cepacia complex. Step 2. Burkholderia multivorans species-specific PCR. By applying primers (BCRBM1 and BCRBM2) that are specific for the species B. multivorans, a single 714-bp amplicon will be generated for strains representative of the species B. multivorans. Step 3. Restriction fragment length polymorphism (RFLP) analysis of the B. cepacia complex recA-gene PCR product. The recA-gene PCR product from step 1 is subjected to digestion with restriction enzyme HaeIII. The generated RFLP profile is going to be compared with the unique profiles generated from reference strains representative of the different species in the B. cepacia complex (49).

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A

B

C

D

Fig. 9.5. Images of 3-day-old P. aeruginosa biofilms grown in flow chambers. Biofilm shown in (a) and (b) are nontreated biofilms. In pictures (c) and (d), biofilms have been treated with tobramycin for 24 h. Live bacterial cells appear green and dead cells appear red.

3.4.1. DNA Extraction

1. One to two colonies of the B. cepacia complex-like bacteria are harvested from an overnight culture on a routine solid medium and thoroughly suspended in 1 ml sterile distilled water. 2. Bacterial suspension (100 μl) is mixed with 130 μl bacteria lysis buffer and 20 μl proteinase K (from the MagNa Pure LC DNA Isolation Kit III) and incubated at 65◦ C for 10 min. The mixture is then incubated at 95◦ C for 10 min to inactivate the proteinase K, followed by cooling for 5 min. 3. The sample is then quick-spin centrifuged for a few seconds. Two hundred and fifty microliters of the supernatant


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is transferred to a well in the sample tray in the MagNa Pure LC machine and the DNA extraction process is started. When the process is finished, the purified DNA product is ready for PCR amplification. 3.4.2. PCR Reaction

1. Purified DNA (10 μl) is mixed with 40 μl PCR mastermix in a microcentrifuge tube. 2. The PCR amplification is carried out using a Flex-Cycler with the following program: a. B. cepacia complex: 94◦ C for 2 min, then 30 cycles of 94◦ C for 30 s, 58◦ C for 45 s, 72◦ C for 60 s, 72◦ C for 10 min. b. B. multivorans: 94◦ C for 2 min, then 30 cycles of 94◦ C for 30 s, 62◦ C for 45 s, 72◦ C for 60 s), 72◦ C for 10 min.

3.4.3. Agarose Gel Electrophoresis

1. PCR product (8 μl) and 2 μl TE EC-1028 (orange) are mixed and the 10 μl mixture is transferred to the application wells in the 3% agarose gel. A molecular marker (100-bp DNA ladder) is also applied. 2. Electrophoretic migration of the PCR product is carried out at 100 V for 45–60 min using TBE buffer containing ethidium bromide. 3. The results are documented by photos using Gel Doc hardware and software (Fig. 9.6, top and middle).

3.4.4. RFLP Analysis

1. PCR product (10 μl) is mixed with 10 μl restriction enzyme mastermix (2 μl NEB2, 0.5 μl HaeIII enzyme, and 7.5 μl distilled water) and incubated at 37◦ C overnight. The restriction enzyme reaction is then stopped with 2 μl EDTA. 2. Cleaved PCR product (10 μl) and 2 μl TE EC-1028 (orange) are mixed and then transferred to the application wells in 10% TBE polyacrylamide gel. A molecular marker (50-bp DNA ladder) is applied in-between every four samples. 3. The electrophoretic separation is carried out at 200 V for 45 min in 1× TBE buffer. 4. After electrophoresis, the polyacrylamide gel is stained in 0.035% ethidium bromide solution for 15–30 min and afterward washed in water for 30 min. 5. The results are documented by photos using Gel Doc hardware and software (Fig. 9.6, bottom). The generated RFLP profile is compared with the unique profiles generated from reference strains representative of the different species in the B. cepacia complex.

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Fig. 9.6. Applying recA-gene PCR-based approach for identification of clinical isolates of the B. cepacia complex from CF patients with chronic lung infection. Top: PCR products after B. cepacia complex-specific PCR. Lanes of 100-bp DNA ladder (MW) and the PCR products of the strains LMG 18830 (Burkholderia cenocepacia), LMG 16654 (B. cenocepacia), LMG 16230 (Burkholderia vietnamiensis), LMG 19182 (Burkholderia ambifaria), LMG 20980 (Burkholderia anthina), LMG 14191 (Burkholderia pyrrocinia), ATCC 25416 (B. cepacia), 10534/08 (clinical strain = B. vietnamiensis), water control (vand), and LMG 18829 (B. cenocepacia). All the isolates generated a specific band at about 1 kbp after the PCR reaction,


3.5. DNA Typing by Pulsed Field Gel Electrophoresis (PFGE) for Typing of P. aeruginosa or Other Species

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1. Overnight peptone broth culture (600 μl) (or any other liquid culture in a shaking flask, 37◦ C) is diluted with 2.4 ml SE buffer (1:5). 2. The suspension is washed with SE buffer (3000 rpm) and resuspended in SE buffer. 3. Sigma agarose (500 μl) and 500 μl resuspended bacteria in SE buffer are mixed at 50◦ C, final agarose concentration is 1%. 4. The agarose blocks are casted in 100-μl tubes sealed with plastic tape and refrigerated (5◦ C) for 15–20 min to become solid. 5. The tape is removed and the agarose blocks are transferred to 1.8-ml round bottom plastic tubes (NUNC) containing ES buffer, and 10 μl proteinase K (50 mg/ml) is added and mixed carefully and left at 56◦ C overnight (15–20 h). 6. In case of “smears” on the PFGE (high endonuclease activity), 250 mg/ml proteinase K is added instead of 50 mg/ml and this procedure is repeated three times on three successive days. 7. The agarose block is removed and the remaining ES buffer and enzyme are discarded. Cold TE buffer (1 ml) is added to the agarose block to stop the enzymatic reaction and left for 1 h. 8. The agarose block is cut into three equally sized pieces (squares) and transferred to a new NUNC tube containing cold TE buffer for 1 h. 9. The TE buffer is removed and 250 μl NE buffer is added and changed three times every 30 min at room temperature. 10. NE buffer 4 (with BSA) containing Spe1 restriction enzyme (150 μl TE buffer + 2 μl Spe1 for one block) is added and left to react overnight at 37◦ C.

Fig. 9.6. (continued) thereby all isolates belong to B. cepacia complex. Middle: B. cepacia complex-specific PCR (left) and B. multivorans PCR (right) results of two clinical isolates (22923/06 and 15782/06). Lanes of 100-bp DNA ladder (MW) and control water (Kontrol vand) and the PCR products. Both isolates generated a specific band at about 1 kbp after the B. cepacia complex PCR, thereby both isolates belong to the B. cepacia complex. The isolate 22923/06 generated a specific band at 714 bp, and thereby the species identification is B. multivorans. The 15782/06 isolate failed to generate a PCR product after the B. multivorans-specific PCR and the species identification has to be further determined by RFLP analysis and it turned out to be a B. cenocepacia. Bottom: RFLP profiles of eight Burkholderia isolates. Three lanes of molecular markers (MW) and HaeIII-cleaved PCR products of ATCC25418 (B. cepacia), LMG 13010 (B. multivorans), LMG 12614 (B. cenocepacia), LMG 14294 (B. stabilis), LMG 16230 (B. vietnamiensis), LMG 19182 (B. ambifaria), LMG 20980 (B. anthina), and LMG 14191 (B. pyrrocinia). Each species presented a unique profile.

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11. The NE buffer with Spe1 enzyme is removed and 1 ml cold TE buffer is added to stop the reaction. It is left for 1 h at room temperature. 12. The TE buffer is removed and 1 ml fresh TE buffer is added. The block is now ready for PFGE. 13. SeaKem agarose (1%) is casted in Tris–borate buffer and the square block is placed at the top of the gel in the appropriate well avoiding air bubbles and the well is sealed with agarose. DNA size markers (Lambda Ladder; BioLabs) are run in triplicate on each gel (left, middle, and right lanes). 14. In case of smear (see step 6), it is necessary to add 500 μM thiourea to the buffer and agarose gel. 15. The PFGE is now run, e.g., on a GenePath PFGE system (Bio-Rad, CA, USA) using Program 1 (PSU) for P. aeruginosa (run time 19.5 h); for other bacteria, shorter or longer time may be required to separate the bands. 16. The PFGE gel is transferred to ethidium bromide (175 μl ethidium bromide (10 mg/ml) in 500 ml Tris– borate buffer diluted 1:20) staining solution and left for 30 min; thereafter it is washed twice for 30 min with distilled sterile water. 17. The PFGE gel is examined and photos are taken, e.g., using the Gel Doc hardware and software for analysis and comparison with other gels stored in the memory (Fig. 9.7). 18. The criterion for evaluating similarity and differences between two gels follows the guidelines by (51, 52), accepting one mutation in a strain leading to disappearance of one lane and appearance of two new lanes, whereas more differences are regarded as different strains. In CF strains during chronic infections, there may sometimes occur clusters of similar but not identical patterns, indicating the evolution by time of the infecting strain. No firm guidelines can be offered in such cases. 3.6. ELISA for Measuring IgG and IgA Antibodies Against Bacterial Sonicate

The Pseudomonas-CF-IgG ELISA kit is a traditional ELISA setup. More than 64 different antigens are detectable in the antigen pool. The results from the pooled human standard serum are used to calculate the concentration of antiserum in the patient sample. Other methods are also available on the market (54).

3.6.1. Principle 3.6.2. Limitations

Non-specific antibodies due to cross-reactivity between P. aeruginosa and other bacterial species are low and correlate with taxonomic relatedness. The human standard antiserum and unknown samples should be assayed in duplicates.


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Fig. 9.7. (Top) Pulsed field gel electrophoresis (PFGE) of molecular weight markers (MW lane) and 12 isolates of colistinresistant P. aeruginosa (lanes 2–13) from 12 CF patients with chronic lung infection and a PAO1 control P. aeruginosa strain (lane K). Restriction enzyme Spe1 was used. Lanes 2, 5, 7, 8, 11, and 13 are identical, and the remaining lanes differ only by 1–2 bands which means that they belong to the same DNA type (55). (Middle) PFGE of molecular markers (3 MW lanes) and 12 isolates of P. aeruginosa (lanes 13–24) from 12 CF patients with chronic lung infection. Lanes 13, 14, 16, 20, and 21 are identical; lanes 15, 17, and 18 differ by more than three lanes (non-identical) and so do lanes 22 and 23 which are identical but differ from lanes 18 and 19 which are identical. (Bottom) From another PFGE gel, a dendrogram has been produced with a computer-based similarity and clustering program (NTSYS; Applied Biostatistics) (29) to facilitate comparison of strains, but this is necessary only if many lanes have to be compared.

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3.6.3. Preparation of Dilutions

Solution A: Antigen solution for coating ELISA plate: Add 100 μl sterile distilled water to the vial containing 9 mg Pseudomonas-CF-IgG antigen and resolve the lyophilized antigen. Dilute the amount to be used the same day 1:2000 in coating buffer. Store the remaining undiluted antigen stock at -20◦ C from where it can be frozen and thawed up to 20 times. Solution B: Dilution of human standard antiserum: Add 100 μl sterile distilled water to the vial with human standard antiserum and resolve the antiserum. Dilute the human standard antiserum to be used the same day by twofold dilution from 1:500 to 1:64,000 in dilution buffer. Store the remaining undiluted human standard antiserum at -20◦ C. The human standard antiserum may be repeatedly frozen and thawed until empty without any change of activity. Solution C: Dilution of patient serum: The patient serum to be measured on the same day has to be diluted 1:100 in dilution buffer (e.g., 10 μl serum added to 990 μl dilution buffer). Solution D: Dilution of rabbit–anti-human IgG HRP: Dilute the rabbit–anti-human IgG HRP 1:20,000 in dilution buffer (e.g., 1 μl rabbit–anti-human IgG HRP is added to 20 ml dilution buffer). Mix thoroughly. Dilute only the amount to be used the same day. 1. Coating of the wells: Add 100 μl diluted antigen (solution A) to each well and incubate for 1 h at room temperature (RT). Aspirate and wash three times with washing buffer. Add 100 μl dilution buffer to each well and incubate for 1 h at RT or overnight at 2–8◦ C. Aspirate and wash two times with washing buffer. 2. Add to each of the appropriate wells either 100 μl of the human standard antiserum dilutions (solution B) or 100 μl diluted patient serum (solution C) and incubate for 1 h at RT. Aspirate and wash three times with washing buffer. 3. Add 100 μl diluted rabbit–anti-human IgG HRP (solution D) to each well and incubate for 1 h at RT. Aspirate and wash five times with washing buffer. 4. Add 100 μl TMB plus standard to each well and incubate for 1 h at RT (dark). 5. Add 100 μl 1 M sulfuric acid to each well and read the absorbance within 10 min using an ELISA reader set to 450 nm.

3.6.4. Calculation of Results

The absorbance of the human standard antiserum dilutions is used to construct a standard curve with OD450 values as a function of log10 ELISA units, e.g.


Dilution factor of human standard serum:

ELISA unit

OD450 value

1:2000

50.00

2.34

1:4000

25.00

1.71

1:8000

12.50

1.13

6.25

0.71

1:16,000

3.6.5. Interpretation of Results

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The normal ELISA unit value of non-infected persons of P. aeruginosa IgG is 0.66 ± 1.64 (mean ± 2 times standard deviation). The 95% upper normal limit is therefore 2.30 and a significant increased titer compared to normal controls is >2.30. The normal ELISA unit value of non-P. aeruginosa-infected CF patients of P. aeruginosa IgG is 0.57 ± 2.39 (mean ± 2 times standard deviation). The normal 95% upper normal limit is therefore 2.96 and a significant increased titer compared to non-P. aeruginosa-infected CF patients is ≥2.96. The difference between non-infected persons and non-P. aeruginosa-infected CF patients is due to crossreactive antibodies induced by, e.g., H. influenzae infections.

Culture positive for P. aeruginosa

Culture negative for P. aeruginosa

IgG ELISA unit > 2.96

A

B

IgG ELISA unit ≤ 2.96

C

D

Group A: Probably chronic P. aeruginosa infection, maintenance therapy is indicated. Group B: Probably not chronic P. aeruginosa infection, but repeat culture. Group C: Probably intermittent P. aeruginosa colonization, eradication therapy is indicated. Group D: Probably not P. aeruginosa colonization or chronic infection. The predictive value of a positive test of Pseudomonas IgG to diagnose chronic P. aeruginosa infection is about 90% and the predictive value of a negative test of Pseudomonas IgG to rule out chronic P. aeruginosa infection is about 90%.

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4. Notes 1. Be aware that NaClO is an alkaline that may cause skin irritation; therefore wear protective clothes, gloves, and goggles. 2. Autofluorescence may appear, make sure that the fluorescent signal is correlated to size and morphology of the investigated cells. 3. It is important that there are no bubbles in the flow chambers. If bubbles are present, try to remove them by gently knocking the flow chamber on the inlet side to the table. Remember to check for bubbles in the flow chamber during the entire experimental period, but after the flow cells are inoculated with bacteria, removing of bubbles should be avoided. If larger bubbles are generated, the biofilm development can be affected, resulting in unusable results. Smaller bobbles can be consumed by the bacteria and will therefore not affect the result. 4. Syto 9 and PI are light sensitive and have to be covered with aluminum foil. Remember to also cover the flow cells with alumina foil during the 15 min of staining. It is possible to add PI to the medium from the beginning of the experiment or when the medium is changed to contain antibiotics and QSI. If this procedure is used, the final concentration of PI in the media has to be 0.0015 mM. Remember to cover every medium containing part with aluminum foil. The flask has to be warmed in a 37◦ C incubator before use. References 1. Westh, J. B. (2001) Pulmonary Physiology and Pathophysiology. Lippincott Williams & Wilkins, Philadelphia, PA. 2. Boucher, R. C. (2004) New concepts of the pathogenesis of cystic fibrosis lung disease. Eur Respir J 23, 146–158. 3. Bjarnsholt, T., Jensen, P. Ø., Fiandaca, M. J., Pedersen, J., Hansen, C. R., Andersen, C. B., et al. (2009) Pseudomonas aeruginosa biofilms in the respiratory tract of cystic fibrosis patients. Pediatr Pulmonol 44, 547–558. 4. Høiby, N., and Frederiksen, B. (2000) Microbiology of cystic fibrosis, in (Hodson, M. E., and Geddes, D. M., eds.), Cystic Fibrosis, 9th ed. Arnold, London, pp. 83–107. 5. Wilson, M. (2005) Microbial Inhabitants of Humans. Cambridge, Cambridge University Press. 6. Høiby, N., and Pressler, T. (2006) Emerging pathogens in cystic fibrosis. Eur Respir

Monog Cyst Fibros Eur Respir Soc 35, 66–78. 7. Skov, M., Koch, C., Reimert, C. M., and Poulsen, L. K. (2000) Diagnosis of allergic bronchopulmonary aspergillosis (ABPA) in cystic fibrosis. Allergy 55, 50–58. 8. Worlitzsch, D., Tarran, R., Ulrich, M., Schwab, U., Cekici, A., Meyer, K. C., et al. (2002) Effects of reduced mucus oxygen concentration in airway Pseudomonas infections of cystic fibrosis patients. J Clin Invest 109, 317–325. 9. Kolpen, M., Hansen, C. R., Bjarnsholt, T., Moser, C., Christensen, L. D., van Gennip, M., et al. (2010) Polymorphonuclear leukocytes consume oxygen in sputum from chronic Pseudomonas aeruginosa pneumonia in cystic fibrosis. Thorax 65, 57–62. 10. Worlitzsch, D., Rintelen, C., Bohm, K., Wollschlager, B., Merkel, N., Borneff-Lipp, M., et al. (2009) Antibiotic-resistant obligate


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Chapter 10 Approaches to Study Differentiation and Repair of Human Airway Epithelial Cells Sophie Crespin, Marc Bacchetta, Song Huang, Tecla Dudez, Ludovic Wiszniewski, and Marc Chanson Abstract One of the main functions of the airway mucosa is to maintain a mechanical barrier at the air–surface interface and to protect the respiratory tract from external injuries. Differentiation of human airway epithelial cells (hAECs) to polarized airway mucosa can be reproduced in vitro by culturing the cells on microporous membrane at the air–liquid interface. Here, we describe approaches to study differentiation as well as repair of the hAECs by using a commercially available airway cell culture model called MucilAirTM . Key words: Human airway epithelial cell culture, MucilAirTM , differentiation, wounding techniques, repair, cystic fibrosis.

1. Introduction The respiratory epithelium plays a fundamental role as a line of defense against pathogens. Among other lung diseases, cystic fibrosis (CF) has been associated with a damaged airway mucosa consequently to chronic lung inflammation and with an abnormal repair (1, 2). Various models were developed by researchers aiming to study the behavior of the respiratory epithelium and its repair. Animal models appear to be useful for CF research (see related chapter in this book). Focusing on lung functions and


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repair, several strategies have already been described in the literature using these models (for a review, see (3)). In vivo approaches consist in the inhalation of gases or the intratracheal instillation of drugs. Ex vivo airway epithelial xenograft models have also been developed. In this case, immunocompromised mice were used to receive grafts subcutaneously. Using this approach, Hajj et al. (4) studied the regeneration of human CF airway epithelium and reported a delayed and abnormal re-differentiation. Nonetheless, these animal-based models exhibit some troubles. First of all, the study of mouse lung regeneration in the context of CF brings into question that mice do not exhibit a CF phenotype in the control conditions (without injury). So, even if we may draw conclusions about repair of the normal epithelium, the extrapolation to the CF epithelium based on CFTR knockout mice might be biased. Second, in the xenograft model, grafts are transplanted heterotypically, leading to a complete change of the normal tissue environment. Closer to the native human healthy or pathologic airway epithelium, several groups developed in vitro cultures using hAECs obtained from surgical resection or brushing. Epithelial cells from the airway epithelium are isolated by explant culture or enzymatic dissociation (for review, see (5)). A key concept is the switch from submerged cultures to an air–liquid interface allowing the development of differentiated airway epithelia (6, 7). To this end, inserts with porous membranes in tissue culture wells were used (8). On the same basis of air–liquid interface culture, at least two kinds of models were developed. One is based on the plating of isolated hAECs on collagen IV-coated inserts (9). The culture is usually viable for about 1 month after the beginning of the culture (4). The other model consists in a first step of hAEC proliferation followed by a second step of differentiation (10, 11). These models present the advantage to amplifying a rare material, especially for CF tissues. The MucilAirTM system presents the characteristics of the human airway epithelium with the presence of basal cells, goblet cells, and ciliated cells organized in a pseudostratified epithelium. An advantage of this system is the possibility to keep the airway epithelium differentiated for up to 6–9 months enabling to monitor epithelial repair, a process that lasts days to weeks depending on the extent of the lesion. The aim of this chapter is to first describe the characteristics of a well-differentiated airway epithelium maintained in primary culture. Thus, different protocols concerning the states of differentiation are given: epithelial morphology and expression of specific markers (Ki67, β-tubulin, and connexins). Second, a strategy mimicking an injury in vitro is described. Finally, the last part of this chapter concerns methods to monitor epithelial repair, including measurement of the kinetics of wound closure, cell proliferation, and migration.

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2. Materials 2.1. hAEC Cultures and Media 2.1.1. hAEC Cultures

2.1.2. hAEC Media

2.2. Other Reagents and Solutions

Differentiation and repair of hAECs were evaluated on the commercially available in vitro airway epithelium cell model MucilAirTM (Epithelix, Plan-les-Ouates, Switzerland). The respiratory epithelium is reconstituted from primary hAECs freshly isolated from nasal polyps or from tracheal/bronchial biopsies, according to methods that are extensively described in this book. Briefly, hAECs are seeded onto 33-mm2 Costar Transwell inserts (Costar, ref. number 3470) with transparent microporous membranes (0.4-μm pore). Two days after seeding, hAECs are switched to an air–liquid interface for at least 45 days. This leads to the differentiation of hAECs to a mucociliated pseudostratified airway epithelium that is maintained in a homeostatic state for months (11). The transparent microporous membrane allows direct observation of the cells under a conventional inverted microscope. Moreover, the polyethylene membranes are more resistant than polycarbonate membranes and can withstand mechanical forces. Typically, the medium used to maintain hAEC cultures at the air– liquid interface is a 3:1 mix of DMEM with GlutaMax (Invitrogen, ref. number 31966-021) and F12 (Invitrogen, ref. number 21765-029), supplemented with penicillin–streptomycin (15,000 units and 30 μg/ml, respectively; GIBCO, ref. number 15140148) and amphotericin B (Amimed, ref. number 4-05F00-H). Other commercially available “ready-to-use” media can also be purchased from Epithelix (Plan-les-Ouates, Switzerland), Clonetics Corp. (San Diego, CA), or PromoCell GmbH (Heidelberg, Germany). 1. Buffered NaCl solution: Isotonic saline solution containing 0.9% NaCl supplemented with 10 mM HEPES and 1.25 mM CaCl2 . 2. Dulbecco’s phosphate buffered saline (DPBS) with calcium and magnesium (GIBCO, ref. number 14040). 3. PFA solution: Make 4% paraformaldehyde solution freshly from powder (Sigma, ref. number P6148) in DPBS. Adjust pH at 7.2. 4. Triton X-100 solution: Make 0.3% Triton X-100 (Sigma, ref. number T8787) solution freshly in DPBS. 5. 0.5 M NH4 Cl solution in DPBS. 6. BSA solution: Make 2% bovine serum albumin solution in DPBS freshly.

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7. Lucifer yellow 4% solution: Prepare the Lucifer yellow solution (Sigma, ref. number L0259) in 150 mM LiCl buffered to pH 7.2 with 10 mM HEPES. Keep the solution at 4◦ C in dark and spin down before use to remove any possible aggregates. 2.3. Materials for Wounding

1. A conventional airbrush (Triplex, Gabbert). 2. A source of compressed air. 3. Pressure regulator (0.1 and 0.5 bar). 4. Flexible pipe allowing the connection between the pressure regulator and the airbrush.

2.4. Other Equipments

1. Inverted microscope equipped with fluorescein filters and UV illumination, standing on an anti-vibration table. A micromanipulator is also required to position thin-tip microelectrodes. 2. Borosilicate glass capillaries with an internal filament (for example, Kwik-FilTM 1B120F-4 from World Precision Instruments, Inc.) and a microelectrode puller (typically, we use a vertical Narishige PC-10, Tokyo, Japan). 3. Module allowing the measurement of the transepithelial electrical resistance (for example, EVOMX from World Precision Instruments, Inc.).

3. Methods 3.1. Approaches to Study Differentiation of hAEC

hAECs grown on filters lose their differentiated features. Differentiation can be triggered by exposing the apical surface of hAECs to air (7). Below, we describe approaches to monitor the differentiation of hAECs to a full polarized airway epithelium.

3.1.1. Morphology of the Differentiating Airway Epithelium

The differentiation of hAEC can be evaluated by paraffinembedded sections of the cultures, although the procedure on Transwell inserts is delicate: 1. Fix the hAEC culture with the 4% PFA solution for at least 1 h. 2. Carefully remove the microporous membrane from the insert using a scalpel blade. Avoid tearing/bending of the membrane. 3. Follow a usual protocol of dehydration (succession of baths of ethanol from 70 to 100%, xylol) and embedding in paraffin. Embed the membrane vertically in paraffin. We


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use an automatic embedding machine for treating histological examination (Tissue Processor; Leica Microsystems, TP1020). 4. Make 5-μm-thick sections with a microtome and mount R slides). See Note 1 them on charged slides (as Superfrost for tips and tricks. 5. Rehydrate the samples (succession of baths of xylol, ethanol from 100 to 70%, distilled water) before staining (hemalun– eosin, periodic acid Schiff coloration, Alcian blue, etc.). The morphology of hAECs grown at the air–liquid interface for increasing the amount of time is illustrated in Fig. 10.1 (top panels). With time, hAECs become taller and cilia appear at the apical surface. The presence of basal and mucous cells is also observed. The typical morphology of the respiratory epithelium is maintained on the long term.

Fig. 10.1. Long-term differentiation of MucilAirTM hAEC cultures. Paraformaldehyde-fixed, paraffin-embedded (PFPE) sections of MucilAirTM cultures. hAECs organized as a monolayer during the first week of culture. At this time, a high proliferation rate (Ki-67 staining) is associated with an undifferentiated state (few cells stained for β-tubulin) and a high level of Cx26 expression. After 7 weeks, the hAEC culture exhibits a pseudostratified ciliated epithelium with goblet cells (arrowhead). This differentiation is correlated with a low proliferation rate, a huge amount of β-tubulin staining, and the absence of Cx26 expression. The latter profile is maintained over months (as shown as an example at 14 weeks). Bar, 25 μm.

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3.1.2. Immunohistochemical Detection of Markers of hAEC Differentiation

Typically, differentiation is associated with inhibition of hAEC proliferation and occurrence of neociliogenesis. Several markers of hAEC differentiation can be detected by immunohistochemistry. To this end, immunofluorescence is performed for Ki-67 (a nuclear marker of cell proliferation) and β-tubulin (a component of ciliae). We also use connexin26 (Cx26, a gap junction protein) and Cx43 as key markers of differentiated hAECs (12): 1. Fix the hAEC culture with the PFA solution for 15 min. Wash with DPBS. 2. Permeabilize hAECs with the Triton X-100 solution for 15 min. Wash with DPBS. 3. To avoid non-specific staining, incubate the culture first in the NH4 Cl solution for 15 min followed by the BSA solution for 30 min. 4. Primary and secondary antibodies are incubated for at least 90 min at room temperature. 5. After careful washing, cut the microporous membrane out of the insert using a scalpel blade and mount between slide and coverslip with a photobleaching-preventing mounting R , Clinisciences, ref. number H-1000; medium (Vectashield Aquamount, Thermo Scientific, ref. number 14-390-5). See Note 2 for tips and tricks. Examples of immunostaining for Ki-67, β-tubulin, and Cx26 at different times of culture are shown in Fig. 10.1. At early time after the air–liquid interface has been established, hAEC cultures exhibit mostly a monolayer appearance, high proliferation rate, and absence of ciliae. With longer time at the air–liquid interface, proliferation ceased while differentiation is evidenced by the absence of Ki-67 detection and numerous cilia covering 90% of the epithelial surface after 45 days of culture. Of interest is the marked decrease in the expression of the gap junction proteins Cx26 and Cx43 with differentiation of hAECs (12), as illustrated in Fig. 10.1 for Cx26.

3.1.3. Monitoring Gap Junctional Intercellular Communication

The loss of Cx43 and Cx26 with time at the air–liquid interface allows evaluating hAEC differentiation by monitoring gap junctional intercellular communication. In human, 20 different genes coding for connexin have been found. These connexins are associated with specific pattern of tissue expression, and depending on connexin composition, gap junction channels exhibit different permeability to molecules and dyes. We describe below an approach to microinject the fluorescent dye Lucifer yellow in hAEC cultures: 1. Cut the top of the Transwell insert half a centimeter above the cell culture. See Note 3 for tips and tricks.


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2. Place the insert on a drop of culture medium onto a glass slide and move the preparation to the stage of the inverted microscope equipped for fluorescein detection. 3. Pull a microelectrode (typically 20–50 M). Bend the pipette over a thin flame (away for the tip) to an angle of about 10–20◦ . This will help to position the electrode onto the cell surface within the narrow space provided by the insert. 4. Fill the pipette tip with the Lucifer yellow solution. See Note 4 for tips and tricks. 5. Connect the microelectrode to the micromanipulator and bring the tip in close contact to the cell surface. The fluorescence of Lucifer yellow helps to locate the microelectrode. The cell impalement should be as gentle as possible to maintain cell viability; usually one brief finger tap on the anti-vibration table is sufficient. 6. Allow the dye to diffuse out of the electrode into the cells for 3 min and then rapidly remove the pipette. Dye coupling can be evaluated immediately by counting the number of fluorescent cells. Use a new electrode for each microinjection. An extensive cell-to-cell diffusion of the tracer will be indicative for a not yet differentiated airway epithelium. Ciliated cells within a well-polarized and differentiated airway epithelium do not communicate in terms of Lucifer yellow diffusion. This does not mean that ciliated cells are devoid of gap junctions; in fact, they express Cx30, another gap junction protein which is not permeable to Lucifer yellow (12). 3.1.4. Cytokine and Mucin Production

The production of IL-8 and mucin is changing with hAEC differentiation. Il-8 was measured using an ELISA kit (CLB, Amsterdam, The Netherlands or BD OptEIATM , BD Biosciences, UK) in basal medium that was collected every 2 days. Typically, a well-polarized and differentiated airway epithelium produces 5–15 ng/ml of IL-8 per day. Quantification of mucin production can be evaluated with the enzyme-linked lectin assay (ELLA). At time 0, the apical surface is rinsed with the buffered NaCl solution and cultures are returned to the incubator for various amount of time. At appropriate time points, the accumulated mucus at the apical surface is recovered with 200 μl of buffered NaCl. Glycoproteins in the mucus are captured by Helix pomatia lectin (HPA–lectin) and then revealed by HPA–horseradish peroxidase lectin conjugate (HPA–HRP). The amount of mucus secreted on the epithelial surface is calculated by dividing the obtained values with the number of days of accumulation (ng/ml/day). Mucins are not present in the basolateral compartment.

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3.2. Approaches to Study Repair of hAECs

Following injury, hAECs migrate and proliferate to cover the wound. Covered areas exhibit hAECs that stop proliferating but re-differentiate into a pseudostratified airway epithelium (13). All these processes take place simultaneously with progression of the wound closure. Below, we describe approaches to wound the airway epithelium and to monitor its repair.

3.2.1. Wounding

Several approaches may be considered for wounding a MucilAirTM culture (see Notes 5–7 for alternative wounding techniques). In the specific context of in vitro hAEC cultures on Transwell inserts, these approaches are limited by the fragility of the microporous membrane and the lack of accessibility to the cell surface. We report below the method we are using to make reproducible, regular, and circular wounds. Targeted cells are locally removed from the insert without damaging adjacent cells on the membrane by using an airbrush linked to a pressure regulator. As shown in Fig. 10.2, the diameter of the airbrush fits in the

Fig. 10.2. Procedure for wounding well-differentiated airway epithelia. a Classic airbrush use for making a wound in hAEC cultures (top). The airbrush and the Transwell insert exhibit nearly similar diameters (middle), allowing standardization of the wound by keeping a distance of 4 mm between the airbrush nozzle and the microporous membrane (bottom). b Typical view of a wound (bottom) made with the airbrush under a 0.5 bar pressure maintained for 1 s. A non-wounded hAEC culture of at least 6 weeks old is shown for comparison (top).


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Transwell inserts (Fig. 10.2a). As these inserts are particularly calibrated, the same distance between the airbrush nozzle and the epithelial surface is kept. Note that these parameters depend mainly on the companies where airbrush and inserts are from: 1. Rinse the apical surface of hAEC cultures with the buffered NaCl solution. 2. Every pieces of the airbrush should be carefully cleaned. Also manage a clean area under sterile conditions (typically a culture hood) for wounding. 3. To avoid drying of the apical surface while wounding, the reservoir of the airbrush is filled with the buffered NaCl solution. 4. Adjust the air pressure between 0.1 and 0.5 bar. 5. Introduce the airbrush head into the insert until the end (about 4 mm). 6. Apply a brief pulse of air for 1–2 s. This parameter is highly dependent on the equipment and should be adjusted by each user. Depending on the pressure and the number of pushes on the airbrush, the size of the wound varies between 2 and 15 mm2 . 7. Remove debris and detached cells by carefully washing the surface with the buffered NaCl solution and return the cultures to the incubator to allow cell repair. Images of the circular wound that is obtained following this procedure are shown in Fig. 10.2b. 3.2.2. Monitoring Wound Closure: Kinetics

One of the most convenient, reliable, and non-destructive methods to monitor wound closure is the measurement of transepithelial electrical resistance (TEER). The TEER of wounded cultures should be compared with that of empty Transwell inserts with culture medium in both basal and apical compartments. With repair and re-establishment of junctional complexes, TEER sharply increases to about 400–500 cm2 , which is typical for in vitro human airway epithelia (see Discussion in reference 11). The kinetics of recovery obviously depends on the wound size. Another approach to monitor the kinetics of wound closure is by image analysis. To this end, we use an automated inverted microscope (DMIRE2; LEICA) equipped with a DMSTC XY stage and a digital camera (Ds-5Mc; Nikon) connected to a personal computer. At regular intervals, the surface area of each insert is scanned. Typically, 35 images using a 5× objective are needed to complete the scanning of one insert. Reconstitution of the culture surface is performed by the analysis of pictures with the Image Pro Plus 6.0 software (Media Cybernetics). Alternatively, the ImageJ software (National Institutes of Health, Bethesda) can also be used (http://rsb.info.nih.gov/ij/). Typical images of progression of hAEC repair are shown in Fig. 10.3a, b also

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Fig. 10.3. Monitoring wound closure. a The wound closure is monitored by the acquisition of images taken at different times using an automated inverted microscope equipped with an automatized XY stage and a digital camera. The area covered within a determined period of time allows estimating the rate of epithelial repair. b View of the morphology of the repairing airway epithelium, as investigated using PFPE sections. Bar, 50 μm.

shows a paraffin section of an airway culture 96 h after wounding. Note the change in the height of the epithelium characterized by non-ciliated migrating/proliferating hAECs. To quantify wound closure, the wound area X is measured at different times (T1 , as a circle, allowing the T2 , etc.). The wound area X is considered √ determination of its radius R:R = (X /π). The distance covered by the migrating/proliferating hAECs between time intervals is then given by the difference between R1 and R2 , with R1 and R2 being the radii determined at times T1 and T2 , respectively. 3.2.3. Monitoring Wound Closure: Cell Behavior

It is also possible to monitor wound healing by live imaging. The approach is limited by many factors, including time (it takes days for the epithelium to repair), cell focus (change from a tall differentiated epithelium to a migrating cell monolayer), maintenance of pH of the culture medium, and humidity of the cell environment. However, live imaging can be used within shorter time frames (several hours) at higher resolution to monitor cell behavior at edges of the wound. Thus, depending on the time after wounding, it is in principle possible to observe cell migration, division, and/or differentiation in specific and restricted areas of the repairing airway epithelium. Typically, one needs an inverted microscope equipped with a close chamber enabling control of temperature, CO2 , and humidity. The microscope is


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also equipped for time-lapse imaging. For good (but not optimal) resolution of cell behavior cultured on Transwell inserts, refer to Note 8 for tips and tricks. 3.2.4. Criteria of Repaired Airway Epithelium

As expected from a well-differentiated and polarized airway epithelium, the very same criteria hold for a fully repaired epithelium after injury, including the following: 1. A tall pseudostratified mucociliated morphology. 2. Positive and massive immunostaining for β-tubulin but no detection of Ki-67, Cx26, and Cx43. 3. Lack of Lucifer yellow-mediated gap junctional intercellular communication. 4. Low basal production of IL-8 but sustained apical secretion of mucins.

4. Notes R 1. Superfrost slides are positively charged allowing a better adhesion of samples and avoiding the use of fixation or glue. Alternatively, regular slides may be used after a poly-lysine treatment.

2. To avoid pressing the airway mucosa during the mounting of the samples after immunostaining, an alternative is to use small strips of tape on each edge of the Superfrost slide to create a small chamber where the sample is placed. Then, cover the sample with a coverslip. 3. To easily cut off the top part of inserts, we used a thin inox wire connected to a 6.3-A, 2–8-V DC generator (typically generators used for microscope bulbs). We designed a system whereby the positioning of the wire height could be changed to adapt for various plastic wares (Transwell inserts, Petri dishes, etc.). 4. To easily backfill microelectrodes with the Lucifer yellow solution, we recommend the MicroFilTM needle from World Precision Instruments, Inc. (Microfil MF34G). 5. Mechanical wound may be performed by using the tip of a pipette (14, 15). This technique is particularly simple when performed on a plastic dish but becomes more challenging on membrane inserts. The main inconvenience remains the risk to damage the microporous membrane and to tear the tall airway epithelium apart. 6. Mechanical wound may also be performed following the methodology reported by Vermeer et al. (16). A

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home-designed wounding device performs a reproducible ring-shaped wound when lowered onto cells and turned 360◦ C. 7. Chemical wounding consists of the deposit of a small volume of 1 M sodium hydroxide and its rapid neutralization with DPBS. Tournier et al. (17) reported that a drop of 1 μl leads to a circular wound area of 30 mm2 . Transposed to our model (insert area of 33 mm2 ), this technique would need to be adapted for smaller volumes. 8. To monitor the cell behavior by live imaging, we are using 35-mm Petri dishes with a glass bottom (12 mm, ref. number 73911035; Milian) and for which a hole has been drilled through the covers. The diameter of the hole is slightly larger than that of the bottom of the insert. This simple trick allows holding the insert in proximity to the glass bottom, which is of help for imaging using a 40× objective. The dish is filled with culture medium, while 100 μl is added to the wounded surface. This allows to bath cells with sufficient medium and to ensure humidity and gas exchange in the Petri dish.

Acknowledgments This work was supported by the grants from the Swiss National Science Foundation and by Vaincre la Mucoviscidose. References 1. Puchelle, E., and Zahm, J. M. (2006) Repair process of the airway epithelium, in (Lenfant, C., and Dekker, M., eds.), Airway Environment: From Injury to Repair. Series Lung biology in health and diseases. Marcel Dekker, New York, NY, pp. 1576–1582. 2. Voynow, J. A., Fischer, B. M., Roberts, B. C., and Proia, A. D. (2005) Basal-like cells constitute the proliferating cell population in cystic fibrosis airways. Am J Respir Crit Care Med 172, 1013–1038. 3. Liu, X., Driskell, R. R., and Engelhardt, J. F. (2006) Stem cells in the lung. Methods Enzymol 419, 285–321. 4. Hajj, R., Lesimple, P., Nawrocki-Raby, B., Birembaut, P., Puchelle, E., and Coraux, C. (2007) Human airway surface epithelial regeneration is delayed and abnormal in cystic fibrosis. J Pathol 211 3, 340–350. 5. Gruenert, D. C., Finkbeiner, W. E., and Widdicombe, J. H. (1995) Culture and transfor-

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differentiated human airway epithelia. Methods for establishing primary cultures. Methods Mol Biol 188, 115–137. Fulcher, M. L., Gabriel, S., Burns, K. A., Yankaskas, J. R., and Randell, S. H. (2005) Well-Differentiated Human Airway Epithelial Cell Cultures. Human Cell Culture Protocols, 2nd ed, Methods Mol. Med., vol 107. Springer, New York, NY, pp. 183–206. Wiszniewski, L., Jornot, L., Dudez, T., Pagano, A., Rochat, T., Lacroix, J. S., et al. (2006) Long-term cultures of polarized airway epithelial cells from patients with cystic fibrosis. Am J Respir Cell Mol Biol 34, 39–48. Wiszniewski, L., Sanz, J., Scerri, I., Gasparotto, E., Dudez, T., Lacroix, J. S., et al. (2007) Functional expression of connexin30 and connexin31 in the polarized human airway epithelium. Differentiation 75, 382–392. Puchelle, E., Zahm, J. M., Tournier, J. M., and Coraux, C. (2006) Airway epithelial repair, regeneration, and remodeling after injury in chronic obstructive pulmonary disease. Proc Am Thorac Soc 3, 726–733.

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14. Planus, E., Galiacy, S., Matthay, M., Laurent, V., Galvrilovic, J., Murphy, G., et al. (1999) Role of collagenase in mediating in vitro alveolar epithelial wound repair. J Cell Sci 112, 243–252. 15. Lechapt-Zalcman, E., Prulière-Escabasse, V., Advenier, D., Galiacy, S., Charrière-Bertrand, C., Coste, A., et al. (2006) Transforming growth factor-beta1 increases airway wound repair via MMP-2 upregulation: a new pathway for epithelial wound repair?. Am J Physiol Lung Cell Mol Physiol 290, L1277–L1282. 16. Vermeer, P. D., Einwalter, L. A., Moninger, T. O., Rokhlina, T., Kern, J. A., Zabner, J., et al. (2003) Segregation of receptor and ligand regulates activation of epithelial growth factor receptor. Nature 422, 322–326. 17. Tournier, J. M., Maouche, K., Coraux, C., Zahm, J. M., Cloëz-Tayarani, I., NawrockiRaby, B., Bonnomet, A., et al. (2006) alpha3alpha5beta2-Nicotinic acetylcholine receptor contributes to the wound repair of the respiratory epithelium by modulating intracellular calcium in migrating cells. Am J Pathol 168, 55–68.

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Section II Omic Approaches to Study Cystic Fibrosis

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Chapter 11 Introduction to Section II: Omics in the Biology of Cystic Fibrosis William E. Balch Abstract Cystic fibrosis (CF) is a disease that manifests itself in the context of cell, tissue, and organismal (patho)physiology. While a strong focus on the cystic fibrosis transmembrane conductance regulator (CFTR) since its discovery in 1989 has dominated the field with a wealth of experiments that have provided substantial insight into protein function and structure, a largely untapped area of high relevance to both our basic understanding of CFTR function and its role in clinical disease is the realization that CFTR operates in the context of a cellular network. This is a composite of protein–protein interactions and specific cellular and subcellular environments that balance ion conductance at the cell surface with trafficking through the exocytic and endocytic pathways to promote tissue hydration. To address challenges critical for understanding the system responsible for CFTR physiology and CF pathophysiology, a new era of technologies and methodologies focused on systems-level approaches to analysis of cell and tissue function has emerged. These technologies focus our understanding on the environment supporting protein function (referred to genomics) and the protein composition of the cell (referred to as proteomics) that dictates function. In this section, four chapters focus on emerging “omic” approaches to understanding the cellular environment imposed by message levels in the cell (genomics), the protein composition of the cell and network of interactions dictating cell and CFTR function (proteomics), and the lipid environment (metabolomics) that dictates the functionality of numerous membrane environments in the cell that are integral to CFTR function. Key words: CFTR, cellular networks, protein–protein interactions, proteomics, subcellular environments, systems biology.

Cystic fibrosis (CF) is a disease that manifests itself in the context of cell, tissue, and human (patho)physiology. A strong focus on the cystic fibrosis transmembrane conductance regulator (CFTR) since its discovery in 1989 has dominated the field with a wealth of key experiments that have provided substantial insight into CFTR function and structure. In contrast, a largely untapped area of high relevance to our basic understanding of both CFTR


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function and its role in clinical disease is the realization that CFTR operates in the context of a cellular network; this is a composite of protein–protein interactions and specific cellular and subcellular proteomic and metabolomic environments that balance ion conductance at the cell surface with trafficking through the exocytic and endocytic pathways to promote tissue hydration. While it is clear from Mendelian genetics that CFTR variants are defective in function, the underlying loss of function(s) and gain of toxic function(s) observed in human lung, intestine, and pancreatic physiologies as a consequence of disconnecting CFTR from its interacting partners in the network are largely unknown. The challenge defined by the network biology that dictates normal and pathologic CFTR function is now recognized as a problem in “omics” biology. Omics biology refers to the “system” or systems biology in a given cell type that is required to generate, maintain, and protect the proteome in health and that is subject to dysfunction leading to human diseases such as CF. To address the challenges critical for understanding the network of interactions responsible for CFTR physiology and CF pathophysiology, a new era of technologies and methodologies focused on systems-level approaches to analysis of cell and tissue function has emerged. These technologies focus our understanding on the environment-supporting protein function (referred to as genomics/transcriptomics) and the protein composition of the cell (referred to as proteomics) that dictates function. At the root of both genomics and proteomics lies metabolomics, that is, the small molecule lipid and soluble metabolite pools of the cell that regulate organismal health in response to nutrition and multiple environmental factors that stress the cell and challenge both the genome and the proteome function. The pathophysiology of CF is intimately connected to each of these omic categories and an understanding of their individual contributions will undoubtedly provide critical insight necessary to provide benefit to clinical disease. In this section, four chapters focus on emerging omic approaches to understanding the cellular environment imposed by message levels in the cell (genomics/transcriptomics), the protein composition of the cell and network of interactions dictating cell and CFTR function (proteomics), and the lipid environment (metabolomics) that dictates the functionality of numerous membrane environments in the cell that are integral to CFTR function. Chapter 12 discusses current technologies applicable to defining the genomic/transcriptomic environments that may contribute to disease and that could serve as hallmarks for disease progression given the ease at which these approaches can be used in the clinic. Chapters 13, 14, 15, and 16 focus on various aspects of defining the proteomics of CFTR function in health and disease. Chapter 14 gives a broad overview of rapidly evolving mass

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spectrometry approaches that can be used to ascertain the precise composition of cellular proteomes, protein complexes responsible for CFTR function, and metabolites that may contribute to function in a healthy cell and which become defective in disease. This chapter is designed to motivate CF investigators to begin to adapt these diverse technologies in their own studies. Chapter 15 (functional genomics) and 16 (lipidomics) give specific detailed methodologies that illustrate application of proteomic approaches and mass spectrometry to the study of CF. Omics coupled to bioinformatics remains an emerging field that tackles the challenging problem of the systems biology of disease, rather than simply focusing on a single target protein, in this case CFTR. The real problem in CF disease affecting human health lies in the disruption of the operation of the system, not just the protein. While we have much to learn, the promise of understanding the system at genomic, proteomic, and metabolomic levels offers the opportunity for future therapeutic approaches to use the system, that is, the biology, to correct the pathobiology of CF disease.

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Chapter 12 Microarray mRNA Expression Profiling to Study Cystic Fibrosis Shyam Ramachandran, Luka A. Clarke, Todd E. Scheetz, Margarida D. Amaral, and Paul B. McCray Jr. Abstract To understand the links between CFTR mutations and the development of cystic fibrosis (CF) phenotypes, it is imperative to study the transcriptome in affected cell types. Microarray expression profiling provides a platform to study global gene expression in detail. This approach may provide the necessary information to segregate phenotypic characteristics of CF, differentiate between genetic or environmental factors, and assess the advent and progression of disease phenotypes. Moreover, if a “CF signature” of genes with altered expression is defined, this can be used to monitor effectiveness of treatment. We provide here detailed protocols and tips for collecting and preserving tissues and cells, and preparing total RNA. We also outline novel strategies for experimental design and data analysis, and describe some powerful gene and pathway discovery tools. Key words: GeneChip, mRNA, hybridization, Gene set enrichment analysis, Ingenuity pathway analysis, ANOVA, GeneGo, normalization, quality control, differential expression.

1. Introduction 1.1. Overview of Microarray Technology in Studying Cystic Fibrosis

Microarrays or nucleic acid arrays are comprised of thousands of nucleic acid fragments immobilized onto a solid substrate (1–3). Such an “array” or a chip can represent the entire genome of a species, the transcriptome of a specific tissue, or may be designed to fulfill a specific function like assaying for splice junctions, SNPs, copy number polymorphisms, or insertions and deletions. In this chapter, we focus on the use of microarrays to generate an mRNA expression profile of a given sample from distinct sources. Thus,


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a well-designed experimental approach in combination with the right microarray chip will be invaluable in studying, for example, the following: • differences in gene expression between cystic fibrosis (CF) and non-CF patients or animal models in a tissue- or a celltype-specific manner; • changes in mRNA expression profile of a tissue or a cell type in response to an intervention or as CF disease progresses; • genotype/phenotype correlations; • effects of modifier genomic loci; • microbiome of the CF or non-CF airway; and • effectiveness of a treatment in correcting the basic defect in CF. This chapter takes the reader through all the steps involved in conducting a successful microarray experiment. While for every technique there are alternatives, the authors recommend kits and protocols that have worked repeatedly and provided reproducible results. Furthermore, this chapter is written from the perspective of using an Affymetrix GeneChip platform (Affymetrix, Inc.) and thus recommends following the manufacturer’s protocol where applicable. The authors also reference a number of published studies that have used microarray technology to ask a variety of questions pertaining to the field of CF (4–17). These mRNA expression profile studies have been published using samples from model cell lines, primary airway epithelial cell cultures, animal models, and human samples (4–17). These references are meant purely as a guide, to help investigators design appropriate experiments depending on the question they wish to address. To date, the field has not seen meta-analysis approaches where all existing microarray data are compared. To achieve this end a variety of tools are required to normalize, compare, and mine these diverse data sets, which will be addressed (18, 19). Although beyond the scope of this chapter, investigators are encouraged to consider the option of RNA deep sequencing (RNA-seq) as an alternative to microarrays. A benefit of deep sequencing is the absence of hybridization bias that is associated with microarrays, allowing the investigator to “assay all RNA species” instead of just the probes designed for the array. A variety of platforms are now available (e.g., Illumina Genome Analyzer, Illumina HiSeq 2000 Roche FLX 454 Genome Sequencer, ABI SOLiD) that can be adapted to generate the mRNA expression profiles from samples (20, 21). Total RNA from samples can be processed using commercially available kits to create a qualitative and semi-quantitative expression profile (see Note 10). Deep sequencing is a rapidly emerging technology that is receiving increasing applications as the associated costs decline.

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2. Materials 2.1. General Requirements

1. –20 and –80◦ C freezers, ice, RNase-free water, RNaseZAP (Ambion). 2. RNase-free 1.5- and 2.0-ml tubes.

2.2. Collection and Long-Term Storage of Tissues and Cells for Microarray Analysis

1. RNAlater RNA Stabilization Reagent (Ambion). 2. RNAlater-ICE RNA Stabilizing Reagent (Ambion) (for liquid nitrogen flash-frozen samples).

2.3. Preparation of Samples for RNA Isolation 2.3.1. Mortar and Pestle/Liquid Nitrogen Method

1. Mortar, pestle, stainless steel spatula, liquid nitrogen, RNaseZAP (Ambion).

2.3.2. Tissue Homogenization

1. Motorized tissue homogenizer, double-distilled water, 100% ethanol, 25% ethanol, RNaseZAP, RNase-free water, 12 mm × 75-mm glass tubes.

2.4. RNA Isolation 2.4.1. TRIzol/TRI Reagent Solution

2.4.2. mirVana miRNA Isolation Kit (Ambion)

2.5. RNA Quality and Optional Cleanup

1. TRIzol reagent (Invitrogen) or TRI reagent (Ambion), refrigerated centrifuge, chloroform, isopropanol, 75% ethanol. 1. Pre-heated (95◦ C) nuclease-free water, sterile RNase-free blades, 5-ml RNase-free tubes. 1. Agilent Model 2100 Bioanalyzer (Agilent Technologies). 2. Qiagen RNeasy Mini Kit (Qiagen). 3. RNase-free DNase kit (Any manufacturer of choice).

2.6. RNA Storage

1. Glycogen, 3 M NaOAc (pH 5.2), pre-chilled (–20◦ C) 100% ethanol (to precipitate RNA).

2.7. Microarray Experiment 2.7.1. Affymetrix Sample Preparation and Microarray Analysis

1. Affymetrix GeneChip one-cycle (Affymetrix, Inc., Santa Clara, CA).

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2. Affymetrix GeneChip (as per user requirement; Affymetrix). 3. Affymetrix Model 450 Fluidics Station. 4. Affymetrix Model 3000 Scanner.

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2.7.2. NuGEN Pico/Exon Method Procedure

1. NuGEN WT-Ovation Pico RNA Amplification System, v.1.0 (NuGEN Technologies, Inc.). 2. WT-Ovation Exon Module, v.1.0 (NuGEN). 3. Qiagen Min Elute Kit (Qiagen). 4. FL-Ovation cDNA Biotin Module V2 kit (NuGEN). 5. Streptavidin–phycoerythrin stain (Molecular Probes, Inc.). 6. Anti-streptavidin antibody (Vector Laboratories). 7. Affymetrix Model 450 Fluidics Station. 8. Affymetrix Model 3000 Scanner. 9. GeneChip Operating Software (GCOS), v.1.4.

2.8. Candidate Gene Discovery Tools 2.8.1. Gene Pattern

Unsupervised Hierarchical Clustering (22).

2.8.2. Gene Set and Pathway Analysis of Candidate Genes

1. Ingenuity pathway analysis (IPA) (23). 2. Gene set enrichment analysis (GSEA) (24). 3. Database for annotation, visualization, and integrated discovery (DAVID) (25). 4. GeneGo MetaCore (26). 5. STRING and related databases (27, 28, 29).

3. Methods 3.1. Collection and Long-Term Storage of Tissue and Cells for Microarray Analysis 3.1.1. RNAlater RNA Stabilization Reagent (Ambion)

(a) It is recommended that tissues be cut into small bits of ∼5 mm3 size, allowing for better preservation of the sample and easy access to tissue later on (see Note 1). (b) Label and fill 2 ml cryo-vials with 500 μl of RNAlater reagent and place not more than 4–5 pieces of tissue per tube. (c) Top off the vial with RNAlater solution and store overnight at 4◦ C. The following morning, store the vials at –80◦ C for long-term storage.


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(d) Primary air–liquid interface (ALI) epithelial cell cultures may also be stored in the following manner. Cut out the membrane containing the cells and drop not more than four filters into a cryo-vial containing 1 ml of RNAlater reagent. (e) To remove frozen tissue for analysis, place the vial in an ice bucket until the RNAlater thaws. Wash the tissue pieces well in sterile RNase-free water. 3.1.2. RNAlater-ICE (Ambion) (see Note 2)

(a) It is recommended that samples be cut into small bits of ∼5 mm3 size before freezing in liquid nitrogen. (b) Soak the frozen tissue in RNAlater-ICE at –20◦ C overnight to allow complete permeation of tissue. Samples may be stored indefinitely at –20◦ C in this state. (c) RNAlater-ICE has a blue dye to help distinguish between RNAlater-ICE-untreated and RNAlater-ICE-treated sample. Thus, it is recommended to wash the sample pieces in sterile RNase-free water before RNA extraction.

3.2. Preparation of Samples for RNA Isolation 3.2.1. Mortar and Pestle/Liquid Nitrogen Method

(a) Clean the mortar, pestle, and spatula. Make sure they are RNase free. (b) Chill the mortar, pestle, and spatula in a dry ice ethanol bath. (c) Remove frozen tissue from –80◦ C. Weigh out tissue to be processed before homogenizing so as to calculate correct reagent amounts for the sample (see Note 3). (d) After weighing, place the tissue in the mortar and add liquid nitrogen. When the liquid nitrogen stops bubbling, the sample is completely frozen. (e) Grind the tissue into a fine powder with the pestle. (f) Transfer the powder to an RNase-free, 1.5-ml tube using the chilled spatula and add lysis/binding buffer depending on the RNA isolation protocol being used.

3.2.2. RNAlater or RNAlater-ICE Method

(a) Remove tissue from RNA stabilization solution and weigh out the required amount (see Note 3). (b) Rinse tissue in sterile, nuclease-free water (see Note 4).

3.2.3. Tissue Homogenization

(a) Use a handheld tissue homogenizer (Tissue-Tearor, PotterElvehjem tissue grinder) attached to a variable speed 3/8-in. drill. (b) Wash the homogenizer in the following solutions between samples, in the following order: double-distilled

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H2 O, 100% EtOH, 25% EtOH, double-distilled H2 O, RNaseZAP, RNase-free H2 O. (c) Label glass tubes (12 mm × 75 mm), add 1.0 ml homogenization buffer (TRIzol, lysis buffer, etc.) to each tube, and pre-chill tubes in ice. (d) Transfer required tissue sample from RNAlater or RNAlater-ICE into glass tube containing homogenization buffer. Recommended tissue sample size: approx. three pieces of 5 mm3 tissue specimens. (e) Tubes containing the tissue and the homogenization buffer must be kept in an ice bucket during the homogenization process. (f) Twelve to 15 passes of the homogenizer through the sample should homogenize the sample completely. Avoid heating the sample (see Note 5). (g) Wash the homogenizer (as in step 1) before using for next sample. 3.3. RNA Isolation 3.3.1. TRIzol/TRI Reagent Solution Methodology

(a) Homogenize sample as described above. Transfer homogenate to a 1.5-ml tube. Add 1 ml TRIzol reagent and mix thoroughly by vortexing. Let the tube sit for 5 min at 4◦ C. (b) Add 200 μl chloroform, shake vigorously for 15 s, let stand for 5 min at room temperature, shake again. Centrifuge at 16,000×g for 15 min at 4◦ C. (c) Transfer upper phase to a new tube, be very careful not to touch or disturb the interface as it contains RNases (see Note 6). (d) Add 500 μl isopropanol, shake. Store for 30 min at –20◦ C. This is a good stopping point and samples may be stored overnight at –80◦ C if required. (e) Centrifuge at 16,000×g for 10 min at 4◦ C. (f) Wash the pellet with ice-cold 75% ethanol, vortex briefly. (g) Centrifuge at 7,500×g for 10 min, remove supernatant, and air-dry the pellet (or use a Speed-Vac). (h) Dissolve the pellet in 50 μl RNase-free H2 O and heat at 65◦ C for 10 min to completely dissolve the RNA. (i) Quantitate the RNA and store at –80◦ C.

3.3.2. mirVana miRNA Isolation Kit (Ambion) Method

Although this kit is sold as a small RNA isolation kit, it has a very effective and reproducible total RNA isolation protocol. We recommend the use of the manufacturer’s protocol to isolate total RNA from both tissues and cells grown on plastic (see Note 7).


3.3.2.1. Variation of Total RNA Isolation Protocol for Primary Cultures

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RNA from primary air–liquid interface (ALI) cultures of airway epithelial cells may be prepared using the mirVANA protocol with the following variation: (a) Using a sterile blade, cut out the filters from culture plate inserts on which the cells are growing. (b) Place a maximum of four filters into a 5-ml tube and add 1.0 ml lysis/binding buffer. (c) Keep on ice for 2 min. Vortex for 30 s. Repeat three times. (d) Transfer all lysis/binding buffer to a fresh 2.0-ml tube. (e) Follow the manufacturer’s total RNA isolation protocol in the mirVANA kit.

3.4. RNA Quality and Optional Cleanup

The authors recommend the DNase treatment of RNA at this stage. The use of a RNase-free DNase kit from any manufacturer will efficiently remove DNA contamination that can confound data in any downstream application. It is recommended to test the quality of RNA isolated on an Agilent Model 2100 Bioanalyzer (Agilent Technologies). Only samples with an RNA integrity number (RIN) over 7.0 should be submitted for microarray analysis. If consistently poor RIN values are achieved, any commercially available RNA cleanup protocol may be used to improve the purity of the sample. A recommended kit is the Qiagen RNeasy Mini Kit (see Note 9).

3.5. RNA Storage (see Note 8)

Short-term storage: RNA dissolved in RNase-free water at –20 or –80◦ C. Long-term storage: RNA may be stored at –20◦ C as ethanol precipitates.

3.6. Considerations in Designing a Microarray Experiment (See Note 11) 3.6.1. Confounding Variables

In order to minimize every possible origin of variation introduced by experimental design, investigators are encouraged to avoid, as far as possible, the following common sources: (1) different lots of reagents including microarrays themselves; (2) different sources or protocols utilized to obtain the experimental samples; (3) different methods in preparing and storing RNA; (4) different people preparing, labeling, and hybridizing the RNA; (5) changes in tissue and RNA integrity of samples over long storage; and (6) performing the hybridizations on different days.

3.6.2. Unknown Confounders

An additional confounding issue that arises with human samples is the potential for differential probe affinity due to variations in the underlying genomic sequence. For example, a single nucleotide

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variation from a G to an A may significantly alter the affinity of several of the eleven 25-nt probes within a probe set. This is not an issue when dealing with a single, highly inbred organism. However, even in highly inbred organisms, experiments that deal with genetic crosses between two strains (e.g., C57BL/6 and DBA2 mice) will have similar probe affinity-based variations. 3.6.3. Batch Effect

There is a practical limit to the capacity of any facility to perform a large number of microarray hybridizations at the same time. If the sample size of the study is large, investigators are recommended to design their experiment to also minimize the confounding variations that arise due to the performance of hybridizations on different days (batch effect). One such strategy is to rearrange the samples such that each batch of hybridizations represents an equal number of samples across all groups being studied. In the case of replicate samples, these should be run on different days to aid in ascertaining the variability between runs.

3.7. Microarray Experiment

Microarray hybridizations are performed by the following two methods at the University of Iowa DNA Core Facility (see Note 12).

3.7.1. Affymetrix Sample Preparation and Microarray Analysis

(a) Only RNA samples that attain a minimum of 7.0 RIN using the Agilent 2100 Bioanalyzer (Agilent Technologies) are processed. (b) Total RNA (5 μg) is processed using the Affymetrix GeneChip one-cycle target labeling kit (Affymetrix) according to the manufacturer’s recommended protocols. (c) The resultant biotinylated cRNA is fragmented and then hybridized to the Affymetrix GeneChip (GeneChip Porcine Genome Array or the GeneChip Human Genome U133 Plus 2.0 array). (d) The arrays are washed, stained, and scanned using the Affymetrix Model 450 Fluidics Station and Affymetrix Model 3000 scanner using the manufacturer’s recommended protocols.

3.7.2. NuGEN Pico/Exon Method Procedure

(a) RNA quality is assessed using the Agilent Model 2100 Bioanalyzer (Agilent Technologies) and a total RNA within the range of ≥500 pg to ≤50 ng is used. (b) The RNA is processed using the NuGEN WT-Ovation Pico RNA Amplification System, v.1.0 along with the WT-Ovation Exon Module, v.1.0 (NuGEN Technologies) according to the manufacturer’s recommended protocols. This generates sense target (ST)-cDNA. (c) After purification of the ST-cDNA yield (Qiagen Min Elute Kit), 5 μg of ST-cDNA is fragmented and labeled with the FL-Ovation cDNA Biotin Module V2 kit (NuGEN


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Technologies) and combined with hybridization control oligomer (B2) and control cRNAs (bioB, bioC, bioD, and creX) in hybridization buffer and hybridized with an Affymetrix GeneChip as per the manufacturer’s recommended protocols. (d) The arrays are washed and stained with streptavidin– phycoerythrin (Molecular Probes). The signal is amplified with an anti-streptavidin antibody (Vector Labs) using the Affymetrix Model 450 Fluidics Station. (e) Arrays are scanned using the Affymetrix Model 3000 (7G) scanner and the data collected using the GeneChip Operating Software (GCOS), v.1.4. 3.8. General Concepts in Data Analysis 3.8.1. Normalization

A critical component in the analysis of microarray data is normalization of the expression array data. Normalization is the process through which the summarized image data from the CEL files are transformed into numerical values that are comparable across arrays. While early strategies are relatively simple (e.g., global scaling), modern protocols use programs such as RMA (30), GCRMA (31), and PLIER (32). These methods normalize all arrays simultaneously, allowing for variability of intensity to be computationally corrected (see Note 13).

3.8.2. Quality Control

As in any experiment, it is important to implement quality control metrics to isolate and remove erroneous data. The most common techniques for microarray data sets include assessment of any embedded control probe sets, assessment of RNA degradation, and the utilization of pattern-finding computational methods to identify outliers. Affymetrix arrays have a set of probe sets to several housekeeping genes (e.g., GAPDH) allowing relative comparison of intensity of the 3 and 5 probe sets. Because 3 -end labeling is used to generate the labeled cRNA, all molecules should hybridize to the 3 probe set and only the handful that generate very long labeled products should hybridize to the 5 probe set. A similar technique may be used to assess overall RNA degradation among all probe sets. With this procedure, each of the 11 probes for every probe set is categorized based upon their relative position (most 5 to most 3 ) in the gene structure. A plot of the average intensity values for each of the 11 categories should have the highest value at the 3 -end. A characteristic dip is observed in the 3 most probe categories in case of RNA degradation. Finally, pattern-finding methods such as PCA (principal component analysis) and hierarchical clustering may be used to identify samples that are drastically different from others of the same class or category.

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3.8.3. Differential Expression

The most common microarray analysis is an assessment of differential expression. This identifies the set of genes that are expressed at significantly different levels. Many methods are available to determine if a gene is differentially expressed. Two such methods grounded in statistics are the student’s t-test and ANOVA (analysis of variance). In general, ANOVA is the more generalized approach, allowing for more than two classes to be analyzed simultaneously and for multiple explanatory variables (if needed). Both methods return a p value, representing the significance of the differential expression. Because genome-wide experiments are characterized by extensive hypothesis testing (one per probe set), approaches designed to compensate for multiple hypothesis testing may be overly conservative (e.g., Bonferroni correction). Therefore, methods such as the false discovery rate (FDR) are recommended (33). Such methods attempt to control the number of expected false positives, i.e., the proportion of genes expected to be incorrectly present in the list of significant (differentially expressed) genes.

3.9. Candidate Gene Discovery Tools

Here, we provide a list of publicly available and proprietary analysis tools that have been selected for their individual merits. These resources come with detailed instructions that are simple to follow and user friendly, but are by no means exhaustive. Thus, the following sections do not provide instructions on how to use these web-based resources but highlight their individual merits and how you may adapt them to your specific needs.

3.9.1. Gene Pattern

Hosted and developed by the Broad Institute, MIT, Gene Pattern is an open-access online analysis resource (22). It is a compilation of modules, each focused at a particular format of analysis. Most useful from the perspective of analyzing microarray data, the cluster analysis and gene set enrichment analysis tools have been widely used by our research groups. Clustering is a method of unsupervised learning, widely used to segregate samples into groups based on their gene expression profiles. The hierarchical clustering tool in Gene Pattern has been used by our group for pattern recognition in microarray data. Hierarchical clustering helps to segregate samples in an unbiased manner into groups based purely on the expression pattern of samples being tested (see Fig. 12.1a). Thus, this tool functions as a first-pass analysis algorithm for investigators to assess if their samples segregate based on genotype, tissue, gender, age, or non-biological confounding factors like batch effects.

3.9.2. Analysis of Variance (ANOVA)

The one-way ANOVA analysis has been used by our group to determine differences in gene expression among independent groups of samples. Typically, the output consists of all genes in


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the microarray experiment, with each gene accompanied by a fold change and a p value (or FDR) to indicate the amount of change and the significance of change in gene expression across the independent groups of samples. Thus, a candidate gene list may be made based on the fold change and significance of differential expression properties of each gene. If candidate gene lists are large, it is recommended to either make the p value more stringent or consider the FDR q value instead of the p value (see Fig. 12.1b,c). Furthermore, cutoffs based on fold change may help narrow down the gene list to the genes that change by the greatest fold. The authors used the Partek Genomics Suite (36) for the ANOVA analysis. 3.9.3. Gene Set and Pathway Analysis of Candidate Genes

While identification of candidate genes is essentially the crux of a well-planned microarray experiment, an investigator is often confronted with the scenario of having either too many candidates or candidates of unknown relevance to CFTR or CF. In this respect, it helps to look at genes from the perspective of their functional relationships based on knowledge of pathways, protein interactions, and sequence composition. A variety of manually curated databases exist online (see below), both open-access and commercial source, that classify gene products based on their functional interactions, cellular localizations, sequence similarities, and binding motifs to name a few. While this information does not necessarily hand the investigator answers on a silver platter, it does fill in several missing links and puts the raw data in a format that may provide more biological meaning to the investigator. Described below are gene discovery tools that have been used by the authors. A word of caution: the following online resources are compiled based on published literature involving microarray studies, immunoprecipitation experiments, etc. Thus, there is a level of inherent error in these analysis tools based on the character of the curation and the interests of the developers, and the investigator is recommended to vigorously validate candidates before studying their biological and functional roles.

3.9.3.1. Ingenuity Pathway Analysis (IPA)

IPA is an extensive, manually curated proprietary database, classifying gene products based on their functional interactions with other gene products (23). The power of IPA lies in its userfriendly approach. Any list of genes can be uploaded in the form of a text file. Herein lies its greatest limitation. The gene list is put together by the investigator, thus the utility of the output is limited by the relevance of the input gene list. IPA, based on its vast database of interactions, then classifies genes into networks, filling in the blanks with transcription factors and other proteins to form complete pathways (see Fig. 12.1c). IPA is extremely user friendly because it allows the user to access network information,

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B

D

Fig. 12.1. The data set used for this analysis consists of primary well-differentiated airway epithelial cultures isolated from seven different normal human donors, grown at an air–liquid interface (ALI) (34). Half of all primary cultures from each donor were subjected to a pro-inflammatory cytokine cocktail treatment comprising IFN-γ, TNF-α, and IL-1β, at a final concentration of 100 ng/ml each for 24 h (35). Total RNA was isolated and cDNA microarray analysis was performed comparing the gene expression profile of these two sample groups. Presented here are data output samples using the following tools: unsupervised hierarchical clustering (Gene Pattern), gene set enrichment analysis (GSEA), ingenuity pathway analysis (IPA), and a heat map of differentially expressed genes (based on ANOVA). (a) Unsupervised hierarchical clustering: Samples 1–, 3–, 6–, 8–, 5–, 7–, and 9– constitute the “untreated” group, while samples 2+, 4+, 10+, 11+, 12+, 13+, and 14+ are treated with the pro-inflammatory cytokine cocktail. The combined unsupervised hierarchical clustering (22) of the 14 samples reveals 2 distinct clusters. All untreated samples segregate and form a separate cluster, while the cytokine-treated samples segregate into a different cluster. The dendrogram height of each cluster reveals that the sample groups have a distinct gene expression profile that is similar within each group but different between the two groups. (b) Heat map of the most significant differentially expressed genes: The one-way ANOVA analysis tool available in the Partek Genomic Suite (36) was used to compare the gene expression profile of the two sample groups from (a) and


205

source literature, and gene product characteristics on site. It also allows the user to modify the presentation to adapt it to the user’s own personal and research requirements. 3.9.3.2. Gene Set Enrichment Analysis (GSEA)

GSEA analyzes microarray data based on a priori defined sets of genes, compiled from published literature describing biochemical pathways and co-expression profiles (24). GSEA stands apart from other analysis algorithms as it looks at pathways and signaling programs as a whole. This enables GSEA to detect whole networks and signaling cascades that might change owing to differences in expression of a few key representative genes of that pathway (see Fig. 12.1d). GSEA draws its strength from the fact that it ranks all genes in the experiment and considers even the smallest change in expression of every gene. Thus, the algorithm compiles the cumulative effect of changes in gene expression within a pathway and determines which gene set has the closest correlation with the phenotype. This complex, three-tier analysis enables the user to view processed data from the perspective of pathways or biological processes and to simultaneously focus on individual genes and their differential expression. Investigators can also develop custom gene sets based on the biology or tissue of their interest and use it with this tool.

Fig. 12.1. (continued) characterize genes based on their differential expression. Change in expression was rated based on false discovery rate (FDR). By setting an FDR stringency of FDR 106 infectious units/mL) (23). Clonally derived cells with the desired properties can be cryopreserved and thawed when needed. After thawing, cells usually retain their vector-producing properties for at least 1–2 months before they need to be replaced. 5. Retroviral/lentiviral gene transfer vectors are available from a number of sources. For constitutive expression of a target gene as well as an antibiotic selection marker, we have had good success using the murine leukemia virus-based Retro-X Q Vectors from Clontech. The pSIREN-RetroQ gene transfer vector is a murine leukemia virus-based vector (Clontech, Cat. #PT3737-5) used for constitutively expressing small hairpin RNA (shRNA). The pSLIK vectors (ATCC, Cat. #MBA-268) are HIV-1-based gene transfer vectors used for inducible shRNA expression (19). On occasion, plasmids purified using the Qiagen methods are contaminated with insoluble material (most likely residue from the column). In such cases, the plasmid DNA is clarified by centrifugation (10,000×g, 5 min), and the DNA in the supernatant is transferred to a new tube.

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6. The pH of the 2× HBS solution is critical for obtaining efficient transfection. We usually make up several 2× HBS solutions that vary in pH between 7.05 and 7.2 and then test to see which 2× HBS solution yields the best vector production. 2× HBS can be stored for several months at room temperature. 7. The selection agent concentrations are based on our experience with passage 0–2 primary hAE cells. Densely growing cells are more resistant to selection agents. Maintaining selection for at least 7 days and passaging dense cells into media with selection agent (if necessary) is recommended. 8. Screening short RNA sequences for effective knockdown is critical. We typically screen four commercially supplied siRNA sequences in cell lines that express the protein of interest (16HBE (13) or UNCN3T (14) cells) followed by quantitative RT-PCR, or Western blot if antibodies are available. We have had good results with the Amaxa cell transfection system (Lonza, Walkersville, MD). Effective sequences (typically 1–2 out of 4 are found) are cloned into the vector of choice using appropriate methods. Screening with siRNAs may not be possible for genes/proteins not expressed in cell lines on plastic (e.g., ciliated cell-specific genes), in which case multiple shRNA vectors and empirical testing on the hAE cells will be necessary to identify active sequences. 9. Appropriate controls are essential, especially for shRNAs potentially having off-target and/or non-target effects (e.g., interferon response). Empty vector, scrambled shRNA, and/or shRNAs to irrelevant genes are good choices. Rescue by expression of an shRNA-resistant point mutant protein is the gold standard (26). 10. Seeding densities: Primary human airway epithelial cells are mortal and require sufficient seeding density. Attachment and growth of cells from different individuals and preparations may vary. Generous seeding densities of passage 0 cells on porous supports (>1.5 × 105 cells/cm2 ) are required to obtain consistent, confluent, well-differentiated ALI cultures. Although it is tempting to expand primary cells on plastic, “overexpansion” should be avoided. Passage 0 cells first grown on plastic dishes should be seeded at not less than 1 × 106 , and preferably 2–6 × 106 , cells per 100-mm collagen-coated dish (or as calculated mathematically for other dish sizes). Under these conditions, the cells should grow to >70% confluence within 5–7 days – if a longer period is required, subsequent growth may be impaired. Cells at >70% confluence, but not >95%

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confluence, should be trypsinized for cryopreservation or subpassage to a porous support or expanded one more round to passage 2 by seeding >1 × 106 cells per 100 mm tissue culture dish. Passage 1 and 2 cells seeded on porous supports at ∼1.5 × 105 cells/cm2 (∼170,000 and ∼0.7 R membranes, × 106 cells per 12- and 24-mm Transwell respectively) should result in confluence within 3–5 days after seeding, at which point an ALI should be established. Lower seeding densities may be fully successful with some specimens, which can be determined empirically with aliquots of frozen cells, but is not possible when plating passage 0 cells, and greater variability is anticipated between different preparations. 11. Production of retroviral/lentiviral vectors from most sources of 293T cells is enhanced up to fivefold by treatment with sodium butyrate. An alternate method of treatment is to leave the sodium butyrate-containing medium with 2% FBS on the cells overnight prior to harvesting the virus the next day. The brief exposure of hAE cells to sodium butyrate in the resulting virus stock does not appear to affect their growth and differentiation properties. 12. Cross-contamination of cell lines is an important concern and it is a good policy to work with only one cell line at a time in the culture hood. 13. The decision to employ a vector system enabling selection is up to the investigator. Selection ensures more uniform and higher level transgene expression in the cell population. However, resistance to selection agents may limit cell downstream utility or options, e.g., in gene expression or knockdown experiments requiring selection. When seeded at the recommended density, the selected cells should be 70–90% confluent by day 6 or 7 but may take longer if the efficiency of infection is low as indicated by abundant cell death occurring during selection. If the cells are not confluent within 10–12 days, they may not be able to differentiate well at an ALI. To achieve adequate infection efficiencies, it is recommended that the minimum titer of retroviral/lentiviral vectors is 2 × 105 infectious units/mL. 14. It is important that ALI cultures are confluent and healthy in order to withstand caprate permeabilization. Access of caprate to the basolateral solution in non-confluent cultures will result in cell exfoliation. 15. We recommend performing pilot experiments with 30 mM sodium caprate exposure to verify that it will not cause excess cell cytotoxicity and cell exfoliation of a given set of wells, and reducing the exposure time if necessary.

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16. Having nearly equivalent viral titers for control and experimental vectors and equivalent survival during selection (determined by cell counting) is an important factor. If cell survival differs greatly, then cell growth and differentiation after subculture to an ALI may be different, causing differences in expression or function independent of the specifically introduced changes, thus confounding the interpretation. As always, replication of studies is necessary to reduce the likelihood of misinterpretation. 17. Although we have demonstrated functional knockdown of ENaC using the pSLIK inducible system, GFP-reporter gene expression using this system was not uniform in different cell types in well-differentiated ALI hAE cultures and was predominantly found in columnar, non-ciliated, non-basal cells. Variations in expression among cells likely result from different levels of construct integration as well as the potential for cell-type-specific preferential expression or silencing. Evidently, the CMV enhancer promoters in the pSLIK vector are silenced in ciliated cells. At this point, further studies are needed to comprehensively determine vector backbone and promoter elements resulting in uniform expression in well-differentiated ALI hAE cells. Thus, initial experiments with the vector system to be employed, using a reporter gene such as GFP, are strongly recommended to determine whether there is appropriate expression in the differentiated cell types of interest. References 1. Lechner, J. F., Haugen, A., McLendon, I. A., and Pettis, E. W. (1982) Clonal growth of normal adult human bronchial epithelial cells in a serum-free medium. In Vitro 18, 633–642. 2. Fulcher, M. L., Gabriel, S., Burns, K. A., Yankaskas, J. R., and Randell, S. H. (2005) Well-differentiated human airway epithelial cell cultures. Methods Mol Med 107, 183–206. 3. Lechner, J. F., and LaVeck, M. A. (1985) A serum-free method for culturing normal human bronchial epithelial cells at clonal density. J Tiss Cult Methods 9, 43–48. 4. Gray, T. E., Guzman, K., Davis, C. W., Abdullah, L. H., and Nettesheim, P. (1996) Mucociliary differentiation of serially passaged normal human tracheobronchial epithelial cells. Am J Respir Cell Mol Biol 14, 104–112. 5. Gazdar, A. F., and Minna, J. D. (1996) NCI series of cell lines: an historical perspective. J Cell Biochem Suppl 24, 1–11.

6. Boers, J. E., Ambergen, A. W., and Thunnissen, F. B. (1998) Number and proliferation of basal and parabasal cells in normal human airway epithelium. Am J Respir Crit Care Med 157, 2000–2006. 7. Stoner, G. D., Katoh, Y., Foidart, J. M., Myers, G. A., and Harris, C. C. (1980) Identification and culture of human bronchial epithelial cells. Methods Cell Biol 21A, 15–35. 8. Lundberg, A. S., Randell, S. H., Stewart, S. A., Elenbaas, B., Hartwell, K. A., Brooks, M. W., et al. (2002) Immortalization and transformation of primary human airway epithelial cells by gene transfer. Oncogene 21, 4577– 4586. 9. Zabner, J., Karp, P., Seiler, M., Phillips, S. L., Mitchell, C. J., Saavedra, M., et al. (2003) Development of cystic fibrosis and noncystic fibrosis airway cell lines. Am J Physiol 284, L844–L854. 10. Gruenert, D. C., Basbaum, C. B., Welsh, M. J., Li, M., Finkbeiner, W. E., and Nadel, J. A. (1988) Characterization of human tracheal

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Randell et al. epithelial cells transformed by an origindefective simian virus 40. Proc Natl Acad Sci USA 85, 5951–5955. Masui, T., Lechner, J. F., Yoakum, G. H., Willey, J. C., and Harris, C. C. (1986) Growth and differentiation of normal and transformed human bronchial epithelial cells. J Cell Physiol Suppl 4, 73–81. Ramirez, R. D., Sheridan, S., Girard, L., Sato, M., Kim, Y., Pollack, J., et al. (2004) Immortalization of human bronchial epithelial cells in the absence of viral oncoproteins. Cancer Res 64, 9027–9034. Gruenert, D. C., Willems, M., Cassiman, J. J., and Frizzell, R. A. (2004) Established cell lines used in cystic fibrosis research. J Cyst Fibros 3 Suppl 2, 191–196. Fulcher, M. L., Gabriel, S. E., Olsen, J. C., Tatreau, J. R., Gentzsch, M., Livanos, E., et al. (2009) Novel human bronchial epithelial cell lines for cystic fibrosis research. Am J Physiol Lung Cell Mol Physiol 296, L82–L91. Coyne, C. B., Kelly, M. M., Boucher, R. C., and Johnson, L. G. (2000) Enhanced epithelial gene transfer by modulation of tight junctions with sodium caprate. Am J Respir Cell Mol Biol 23, 602–609. Wu, Q., Lu, Z., Verghese, M. W., and Randell, S. H. (2005) Airway epithelial cell tolerance to Pseudomonas aeruginosa. Respir Res 6, 26. Luo, J., Deng, Z. L., Luo, X., Tang, N., Song, W. X., Chen, J., et al. (2007) A protocol for rapid generation of recombinant adenoviruses using the AdEasy system. Nat Protoc 2, 1236–1247. Cockrell, A. S., and Kafri, T. (2007) Gene delivery by lentivirus vectors. Mol Biotechnol 36, 184–204. Shin, K. J., Wall, E. A., Zavzavadjian, J. R., Santat, L. A., Liu, J., Hwang, J. I., et al. (2006) A single lentiviral vector platform for microRNA-based conditional RNA interference and coordinated transgene

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expression. Proc Natl Acad Sci USA 103, 13759–13764. Jones, L. C., Wonsetler, R. L., Olsen, J., Davis, W. C., Randell, S. H., Stutts, M., and O’Neal, W. K. (2008) Short hairpin RNA knockdown in well-differentiated human bronchial epithelial cells: A method for querying gene function relevant to cystic fibrosis. Pediatr Pulmonol Suppl 31, 284. [Abstract]. Randell, S. H., Walstad, L., Schwab, U. E., Grubb, B. R., and Yankaskas, J. R. (2001) Isolation and culture of airway epithelial cells from chronically infected human lungs. In Vitro Cell Dev Biol Anim 37, 480–489. Dull, T., Zufferey, R., Kelly, M., Mandel, R.J., Nguyen, M., Trono, D., et al. (1998) A third-generation lentivirus vector with a conditional packaging system. J Virol 72, 8463– 8471. Johnson, L. G., Mewshaw, J. P., Ni, H., Friedmann, T., Boucher, R. C., and Olsen, J. C. (1998) Effect of host modification and age on airway epithelial gene transfer mediated by a murine leukemia virus-derived vector. J Virol 72, 8861–8872. Salmon, P., Oberholzer, J., Occhiodoro, T., Morel, P., Lou, J., and Trono, D. (2000) Reversible immortalization of human primary cells by lentivector-mediated transfer of specific genes. Mol Ther 2, 404–414. Sanlioglu, S., Williams, C. M., Samavati, L., Butler, N. S., Wang, G., and McCray, P. B., Jr. et al. (2001) Lipopolysaccharide induces Rac1-dependent reactive oxygen species formation and coordinates tumor necrosis factor-alpha secretion through IKK regulation of NF-kappa B. J Biol Chem 276, 30188–30198. Cullen, B. R. (2006) Enhancing and confirming the specificity of RNAi experiments. Nat Methods 3, 677–681.

Chapter 19 Comparative Biology of Cystic Fibrosis Animal Models John T. Fisher, Yulong Zhang, and John F. Engelhardt Abstract Animal models of human diseases are critical for dissecting mechanisms of pathophysiology and developing therapies. In the context of cystic fibrosis (CF), mouse models have been the dominant species by which to study CF disease processes in vivo for the past two decades. Although much has been learned through these CF mouse models, limitations in the ability of this species to recapitulate spontaneous lung disease and several other organ abnormalities seen in CF humans have created a need for additional species on which to study CF. To this end, pig and ferret CF models have been generated by somatic cell nuclear transfer and are currently being characterized. These new larger animal models have phenotypes that appear to closely resemble human CF disease seen in newborns, and efforts to characterize their adult phenotypes are ongoing. This chapter will review current knowledge about comparative lung cell biology and cystic fibrosis transmembrane conductance regulator (CFTR) biology among mice, pigs, and ferrets that has implications for CF disease modeling in these species. We will focus on methods used to compare the biology and function of CFTR between these species and their relevance to phenotypes seen in the animal models. These cross-species comparisons and the development of both the pig and the ferret CF models may help elucidate pathophysiologic mechanisms of CF lung disease and lead to new therapeutic approaches. Key words: Lung biology, tracheal xenograft, CFTR processing, pig, ferret, mouse.

1. Introduction Animal models that reproduce the human cystic fibrosis (CF) disease phenotypes are required to effectively develop methods to treat the disease. These models also serve to increase our understanding of disease pathophysiology, cystic fibrosis transmembrane conductance regulator (CFTR) processing and channel function, testing of therapeutic molecules, and development M.D. Amaral, K. Kunzelmann (eds.), Cystic Fibrosis, Methods in Molecular Biology 742, DOI 10.1007/978-1-61779-120-8_19, © Springer Science+Business Media, LLC 2011

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of gene therapy approaches. It would be ideal if a single model was available that completely modeled the human disease (see Table 19.1); however, species differences are clearly apparent in the three CF models generated to date. As has been the case for CF mouse models, differences in the severity of various aspects of CF organ disease in the pig and ferret models will likely inform new biologic discoveries about CFTR functions in organ physiology and how dysfunction of these processes lead to disease in humans. The purpose of this chapter is to briefly review the general phenotypes of the CF mouse, pig, and ferret models, and to provide methods for comparative analysis of CFTR biology between these models. More detailed reviews of CF mouse models have been reported elsewhere (1–5), and the methods of construction of the CF pig and ferret models have also been previously reported (6, 7) and will not be a focus of the chapter. Comparative differences and similarities among these models will greatly enhance our understanding of the disease and accelerate the development of a cure for CF. It is important to emphasize that differences among species in their ability to model CF (see Table 19.2) will likely help to educate the field on what factors influence phenotypic variability seen in CF patients. 1.1. The Murine Models of CF

Murine models of CF have existed since the early 1990s, contributing invaluably to the current understanding of CF. To our knowledge, at least 14 mouse models of CF exist, including null and mutant forms of CFTR (8–21). The degree to which these models recapitulate various organ pathologies seen in human CF disease varies. Generally, the severity of the phenotypes in each of these models varies slightly based on the levels of CFTR mRNA, as a result of the gene targeting method used and the genetic background of the mouse (1, 4, 5). Briefly, most of the models display one or more of the following phenotypes including severe abnormalities in the gastrointestinal tract, failure to thrive, decreased rates of survival due to intestinal complications, and hyperinflammatory responses in the airway. Furthermore, most of these CF mouse models retain defects in cAMP-inducible chloride permeability in the nasal epithelium as seen in humans. Though not extensively studied in each model, reports have suggested decreased mucociliary clearance (22–24), reduced fertility (25, 26), mild pancreatic dysfunction (27–29), and liver abnormalities (28). However, these models thus far lack the development of significant spontaneous lung disease as observed in humans with CF. Furthermore, gut obstruction phenotypes seen in CF mice at weaning are clinically different from meconium ileus seen in newborn CF infants and suggest some level of biologic differences in the developmental control of chloride movement in the gut by CFTR between mice and humans.

1480

1476

1482

1484

∼80 kg

∼25 g

∼90 kg

∼2–3 kg

Human

Mouse

Pig

Ferret

91

92

78

100

a Protein sequence identity compared to the human CFTR sequence

AA, amino acid; SMG, submucosal gland

CFTR AA No.

Avg. mass

Species

CFTR identitya (%)

Table 19.1 General species characteristics of CFTR

42

114

21

280

Days of gestation

8

10

6

1

Avg. litter size

4–6 months

6–8 months

6–8 weeks

10–16 years

Sexual maturation

8–10

10–15

2

∼78

Abundant

Abundant

Rare

Abundant

Avg. life expectancy (years) SMG abundance

Animal Models of CF 313

None

Adult: severe (40)

Newborn: frequent (39) Adult: severe (unpublished)

Mouse

Pig

Ferret

Birth: 100% ADD 0% EPD (39)

Birth: 100% EPD (38, 64)

Birth: none Adult: mild ADDa (4, 61)

Birth: ∼72–90% ADD ∼3% EPD Adult: ∼83% EPD (41, 42, 50)

Exocrine pancreas

Birth: 75% MI 100% MA (39)

Birth: 100% ELFTs (39)

Birth: 20% FBC (38, 64)

Birth: normal Adult: mild FBCa (4,61, 63)

Birth: OB at weaninga Adult: MAa (4,61, 62) Birth: 100% MI (38, 64)

Birth: ∼50% ELFTs Adult: ∼10–20% FBC (56, 57)

Liver

Birth: ∼10–15% MI ∼80% MA Adult: MA, OB (51–55)

Gastrointestinal

Birth: normal (39)

Birth: 100% (38, 64)

Birth: normal

Birth: ∼23% (52, 58)

Micro-gallbladder

CBAVD: present at birth (39)

CBAVD: variable at birth (40, 40a)

Reduced female fertilitya (26)

Male infertility: ∼95% CBAVD (59, 60)

Fertility

MI, meconium ileus; TBD, to be determined; ADD, mild lesions associated with exocrine acinar duct dilatation; EPD, severe lesions associated with exocrine pancreas destruction; ELFTs, elevated liver function tests; FBC, focal biliary cirrhosis; MA, malabsorption; OB, intestinal obstruction; CBAVD, congenital bilateral absence of vas deferens a Observed only on certain background strains and/or CFTR genotypes

Birth: infrequent Adult: severe (49)

Spontaneous lung infections

Human

Species

Table 19.2 Cystic fibrosis phenotype across species

314 Fisher, Zhang, and Engelhardt

Animal Models of CF

315

Although the nasal bioelectric defects seen in the CF mouse models appear to closely resemble those in the human nasal epithelium (5), the bioelectric characteristics in the tracheal airways of mice and humans diverge significantly with murine models demonstrating cAMP-inducible changes in chloride permeability despite the absence of CFTR (30, 31). Potential explanations for this observation include differences in airway cell biology (i.e., different types of secretory cells are found in the proximal airway of humans [goblet cells] and mice [Clara cells]), differences in the distribution of submucosal glands (i.e., throughout the cartilaginous airways in humans and located only to the proximal regions of the trachea in mice), and the presence of alternative non-CFTR chloride channels in mice capable of activation in response to cAMP that are not found in humans (5, 30). Much effort has been placed to develop methods capable of studying bacterial clearance defects in CF mouse lung (2, 4, 32, 33). Many groups with variable success have attempted inoculation of these CF mouse models with bacteria, mainly Pseudomonas aeruginosa, to reproduce the human lung disease phenotype. A variety of inoculation methods have been attempted from aerosolization of free bacteria to insertion of bacteria-laden agar beads. Some studies report decreased survival of the CF mice compared to their littermates, while others indicate no difference in clearance between genotypes (2, 4). However, some groups have reproducibly observed excessive inflammatory response and higher mortality to inoculation of the CF mouse lung with bacteria-laden agar beads, despite no differences in bacterial clearance (32, 33). Another interesting model recently reported by Hodges and colleagues (21, 34) is the development of a conditional CFTR knockout mouse model. This model is being used to direct tissue-specific deletion of CFTR following crossing with transgenic mice that directs Cre recombinase expression under tissuespecific promoters. This system will allow for a systematic and directed evaluation of CFTR function at the level of individual organs. 1.2. New Methods for Generating Larger Animal Models of CF Disease

Given the observed phenotype in CF mouse models, it is clear that additional larger animal models of CF would be of utility to the field. Several parameters influence the choice of alternative species to model CF including (1) the types of cells in the airway in comparison to human (2), the distribution of submucosal glands which are thought to play an important role in CF airway disease (3), conservation of CFTR structure and function (4), the composite of alternative chloride channels in the airway, and (5) the reproductive parameters of the species which will make it feasible to rapidly perform research studies. Generation of larger CF

316


animal models such as pig (35), ferret (36), and sheep (37) has been considered, but the technology to manipulate the genomes of these animals has lagged behind until recently. Recently, methods for generating both pig and ferret CF models have been developed using recombinant adeno-associated virus (rAAV)mediated gene targeting of exon 10 (6, 7). The phenotypes of newborn CFTR knockout pigs and ferrets have also recently been described (38, 39) and will be discussed in more detail below. 1.2.1. The Porcine Models of CF

Generation of the CFTR–/– and F508 alleles was accomplished by rAAV gene targeting of male porcine fetal fibroblasts (7). The targeted nuclei were subsequently used for somatic cell nuclear transfer, and CFTR+/– male piglets were born and bred to homozygosity. Due to the lack of reports on the heterozygous F508 piglets, this section will focus mostly on the CFTR–/– piglets. Rogers et al. (38) recently reported a detailed description of the newborn CFTR–/– piglet phenotype. In summary, these pigs were born with near-Mendelian ratios of 1:2:1 with no differences in the newborn birth weight or in appearance between genotypes. CFTR-deficient piglets lacked CFTR mRNA and therefore expressed no protein. Nasal transepithelial potential difference (TEPD) displayed a lack of cAMP-inducible chloride permeability and an elevated baseline TEPD in the CFTR–/– piglets similar to that seen in CF humans and mice lacking functional CFTR. All of the CFTR-deficient piglets developed meconium ileus (MI) and atretic microcolon distal to the obstruction, resulting in failure to pass stool and gain weight after birth. If untreated by surgery, the MI was lethal in 100% of the CFTR–/– animals. To live beyond the first few days after birth, all animals required an ileostomy bypassing the obstruction. Rogers et al. also reported adipose infiltration of the pancreas and complete exocrine pancreatic insufficiency (or destruction) at birth in all CFTR–/– animals. These CFTR-deficient piglets also presented with focal biliary cirrhosis and developed a mucus- and bile-filled micro-gallbladder. No overt abnormalities were seen in the lungs, airways, submucosal glands, male reproductive tract, and other non-CF-related organs. Encouraging to the CF field was the recent report that aged (>2 months) CFTR–/– and CFTR–/F508 piglets developed a CF-like lung phenotype (40). We anticipate future reports describing the lung phenotype of F508/F508 homozygous pigs.

1.2.2. The Ferret Model of CF

The generation of CFTR null ferrets was described in detail by Sun et al. (6). rAAV was used to introduce a stop codon and a neomycin cassette into exon 10 of the CFTR gene in primary ferret fibroblasts. Infected fibroblasts were cloned and selected by serial dilution into G418 followed by PCR screening for the targeting events. Due to early senescence of gene-targeted primary

Animal Models of CF

317

ferret fibroblasts (not an issue with the generation of the CF pig models), it was necessary to rejuvenate gene-targeted fibroblast clones by somatic cell nuclear transfer. Primary fibroblasts were then expanded from 21-day fetuses and used for a second round of somatic cell nuclear transfer. Using this process, eight CFTR+/– male ferret founders were obtained and expanded to generate CFTR+/– breeder pairs. Similar to the CF piglets, there was no prenatal lethality associated with CFTR deficiency in ferret kits. Newborn CFTRdeficient newborn ferrets (kits) failed to thrive compared to their wild-type and heterozygous littermates. Approximately 75% of the CFTR-deficient kits failed to pass meconium due to MI and died within the first 36–48 h of life due to intestinal perforation and sepsis (39). Interestingly, this variable penetrance of MI seen in the CF kits is significantly different than the CF porcine model in which 100% of piglets presented with MI (see Table 19.2). Interestingly, the CFTR-deficient kits that passed stool (∼25%) also failed to thrive and died within the first week of life, despite the fact that their intestinal tract was grossly and microscopically normal. The reason for death of these animals appeared to be due to malabsorption, demonstrating a histologic depletion of fat stores and progressive decline in blood cholesterol with age that was not corrected by pancreatic enzyme replacement. CFTRdeficient kits histologically demonstrated mild pancreatic disease at birth, presenting with exocrine acinar ducts that were swollen with inspissated secretions. Such findings are more similar to the human CF pancreatic phenotype at birth (41, 42) and contrasted with the CF pig model in which nearly complete exocrine pancreatic destruction was reported (38). CF kits also demonstrated early signs of functional liver disease (as evident by elevated liver function tests) despite liver histology not overtly different from controls. Unlike the CF pig model, CF newborn ferrets had a histologically normal gallbladder and the majority of kits that escaped MI presented with bronchopulmonary pneumonia at the time of death. It is currently unclear if the bronchopulmonary pneumonia seen in nutritionally compromised CF kits was secondary to their compromised health status and the inability to clear aspirated material from the lung. Based on the clinical blood chemistries seen in CFTR-deficient kits, drug and nutritional therapy to enhance fat absorption by the intestine was undertaken and significantly improved weight gain and survival (39). This treatment involved oral gavages with ursodeoxycholic acid (to treat apparent liver disease indicated by elevated liver enzymes in the blood), omeprazole (to raise gastrointestinal track pH), and elemental diet. This treatment regime improved liver function tests and raised serum cholesterol. Despite improved weight gain using these treatments, CF kits still often developed lung infections within the first four weeks of life that where characterized

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predominantly by steptococcus and staphylococcus infections. Rearing animals on antibiotics has also been used to enhance survival during the preweaning period. Although the adult CF ferret lung phenotype remains under investigation, a slow progressive and fatal lung disease has been shown to occur in this model (unpublished data). The finding that CFTR-deficient ferrets are susceptible to lung infections both early and late in life is encouraging. 1.3. Future Directions for Larger CF Animal Models

The creation of ferret and pig CF models will undoubtedly enhance CF research for decades to come. Comparative aspects of disease between CF mouse, ferret, and pig models should enlighten mechanisms of CFTR function and CF pathophysiology responsible for diverse disease phenotypes seen in CF patients. Despite the promise that comes with the new CF ferret and pig models, there remain significant barriers to their widespread use in research. Foremost among these barriers are the severe intestinal phenotypes at birth in both models. In the CF pig, the requirement for surgery to treat MI will significantly impair the use of this model for the average researcher. Similarly, since 75% of CF ferrets also present with a lethal MI phenotype at birth, the cost of implementing this model will be quite high until this problem can be solved. Two approaches are currently under investigation to reduce the severity of MI in these models. In the context of the ferret, the variable penetrance of MI suggests that there may be heritable influences on the severity of early intestinal disease in this model. To this end, the CF ferret model is being bred into different genetic lines in an effort to determine if a colony of CF ferrets with reduced penetrance of MI can be generated. Second, gut-corrected transgenic CFTR–/– ferrets that express ferret CFTR under the rat fatty acid-binding protein (FABP) promoter have been generated and shown to correct MI in newborn CF kits (39). This has proven feasibility that a transgenic complementation approach combined with somatic cell nuclear transfer can further improve both the CF ferret and pig models. This approach has been used successfully in CF mouse models to prevent intestinal obstruction at weaning (12). Future development of the gut-corrected CF ferret model will require regenerating the model on a heterozygous background for breeding expansion. A second intriguing aspect of future ferret and pig models of CF pertains to the ability of a particular species to model certain mutations of CFTR. As discussed in more detail below, analysis of pig F508-CFTR demonstrates that this protein is partially processed to the apical membrane of airway epithelia where it retains some level of function (43, 44). These findings suggest that the pig may not be the best model on which to study the F508CFTR mutation. It is currently unclear if ferret F508-CFTR will retain similar processing defects as seen with the human

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mutant protein and a conclusive answer to this question in a ferret F508-CFTR model is in progress (45). Three methods useful in evaluating CF animal models will be discussed in this chapter. The first area will review methods for evaluating TEPD in tracheal xenografts. The second area will review antibodies that are suitable for studying alternative species of CFTR. The last area will review methods for studying species-specific processing of CFTR by metabolic pulse-chase labeling. We will use ferret as a model for most of the methods discussed, since other species have already been published elsewhere. 1.4. Cross-Species Analysis of Tracheal TEPD in a Tracheal Xenograft Model

Analysis of the bioelectric properties of cAMP-inducible chloride channels in the airway is critical to characterizing the ability of a particular species to model CF. TEPD measurements are an effective method to study chloride channel defects and have been extensively used in human and mouse models to study CF. Typically, this assay has been performed on the nasal epithelium since it is readily accessible in a live host. However, given the fact that the CF mouse retains nasal but not tracheal chloride transport defects seen in CF humans, methods to directly assess the bioelectric properties of the trachea in CF animal models are needed. Tracheal TEPD measurements can be obtained ex vivo in a tracheal xenograft model. Freshly excised tracheas cannulated with flexible plastic tubing and inserted subcutaneously in athymic Nu/Nu mice have allowed analysis of tracheal bioelectric properties in a vascularized airway free from infection. This system has been extremely useful in characterizing TEPD in these models due to the early intestinal complications in the newborn ferret and pig CF models. Perfusion of pharmacological ion channel agonists and antagonists allows for a systematic measurement of the bioelectric properties in these new models. This chapter will focus on the protocol for making tracheal TEPD measurements using this ex vivo system. Xenograft cassette design, implantation of xenografts, and maintenance of the xenografts will not be discussed in detail because these general methods have been detailed in an earlier edition of this book (46).

1.5. Cross-Species Analysis of CFTR Processing

As new CF models are developed with specific CFTR mutations, a clear understanding of comparative CFTR biology is paramount. Topics relevant to modeling CFTR mutations in a new species include how closely each of the following resembles that of the human mutant CFTR : (1) the efficiency of folding, (2) the efficiency of detection by endoplasmic reticulum associated protein degradation (ERAD), (3) the stability at the plasma membrane, and (4) the activity of the channel at plasma membrane. Reports have already shown that F508-CFTR processing differences exist among mice, pigs, and humans (43, 44, 47). CFTR is

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composed of five structural domains, including two membranespanning domains, two nucleotide-binding domains, and a regulatory domain. With the exception of the regulatory domain, the other two domains are highly conserved between most model species and humans, more so in pigs and ferrets than in mice. Differences in the primary protein structure among species may dictate the ability of these species to correctly model the CF disease seen in humans. The mouse, pig, and ferret CFTRs are 77, 92, and 91 identical to the human CFTR, respectively (Table 19.1). The identification of species-specific differences in CFTR processing may also help to inform new approaches to enhance processing of human mutant CFTR, by identifying molecular targets responsible for variation among species (i.e., differences in chaperone interactions). This section will focus on CFTR antibody optimization across species and comparative metabolic pulse-chase experiments.

2. Materials 2.1. TEPD Measurements in a Tracheal Xenograft Model

1. Tracheal xenograft at least 4 weeks post-transplantation (see Note 1). 2. Ketamine (100 mg/kg) and xylazine (20 mg/kg) in PBS. 3. Multi-range, variable-rate infusion pump (Orion Research, Cat. no. 001967). 4. pH/mV meter (Fisher Scientific, Cat. no. 13-636-AB15P). 5. Calomel reference electrodes (Fisher Scientific, Cat. no. 13-620-51). 6. 21-Gauge × 0.75-in. butterfly infusion set (Abbott Laboratories, Cat. no. 4492). 7. Computer with data acquisition software (CyberComm Pro 2.3; Fisher Scientific) for recording PD in millivolts. 8. 10-mL Disposable syringes with 21-gauge × 1.5-in. needles. 9. Manifold pump tubing (PVC Solvent Flexible tubing; Fisher, Cat. no. 14-190-139). 10. Silicone tubing (Bio-Rad Laboratories, Cat. no. 7318211). 11. Agar Nobel (Difco Laboratories, Detroit, MI, Cat. no. 0142-01). 12. 1 M KCl. 13. Hemostat. 14. HEPES phosphate-buffered Ringer’s (HPBR) solution: 10 mM HEPES (pH 7.4), 140 mM NaCl, 5 mM KCl,

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1.2 mM MgSO4 , 1.2 mM Ca gluconate, 2.4 mM K2 HPO4 , and 0.4 mM KH2 PO4 . 15. Chloride-free HPBR solution: 10 mM HEPES (pH 7.4), 140 mM Na gluconate, 5 mM K gluconate, 1.2 mM MgSO4 , 1.2 mM Ca gluconate, 2.4 mM K2 HPO4 , and 0.4 mM KH2 PO4 . 16. Ham’s F12 medium. 17. PD buffer sequence: a. HPBR solution, 100 μM 4,4 -diisothiocyanatostilbene2,2 -disulfonic acid (DIDS). b. HPBR solution, 100 μM DIDS, 100 μM amiloride. c. Chloride-free HPBR solution, 100 μM DIDS, 100 μM amiloride. d. Chloride-free HPBR solution, 100 μM DIDS, 100 μM amiloride, 200 μM 8-cpt-cAMP. e. Chloride-free HPBR solution, 100 μM DIDS, 100 μM amiloride, 200 μM 8-cpt-cAMP, 50 μM CFTRINH GlyH-101. 2.2. Cross-Species Analysis of CFTR Processing Requires Antibodies that Efficiently Bind Across Species

1. Immunoprecipitation. 2. In vitro phosphorylation: a. PKA phosphorylation buffer (per sample): 50 mM KH2 PO4 (pH 6.8), 2 μg BSA, 2 μg protein kinase A (PKA; Calbiochem, La Jolla, CA) diluted to 20 μL using ddH2 O. b. ATP phosphorylation buffer (per sample): 50 mM KH2 PO4 (pH 6.8), 4 μg BSA, 10 mM MgCl2 , and 3.6 μCi (6000 Ci/mmol) [γ–32 P]-ATP (Perkin Elmer, Waltham, MA) diluted to 40 μL using ddH2 O. c. Thermal-controlled shaker.

2.3. Cross-Species Analysis of the Processing Efficiency and Stability of CFTR by [35 S]Methionine Pulse Chase

1. Starvation media, DMEM lacking methionine and cysteine (Invitrogen, Cat. no. 21013024). 2. [35 S]Methionine and [35 S]cysteine EasyTag Express 35 S protein labeling mix (Perkin Elmer, Cat. no. NEG772007MC). 3. Activated charcoal-loaded syringe. 4. Chase media, DMEM containing 10% FBS, 1% penicillin and streptomycin, and 2 mM cold methionine and cysteine. 5. Ice-cold PBS. 6. RIPA buffer (150 mM NaCl, 20 mM Tris–HCl, 1% Triton X-100, 0.1% SDS, 0.5% deoxycholate, pH 8.0) containing protease inhibitors (Roche, Cat. no. 34342). 7. Refrigerated tabletop centrifuge.

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8. 1.5- and 2.0-mL Eppendorf tubes. 9. Anti-CFTR antibodies M3A7 and MM13-4 (Millipore, Cat. nos. 05-581 and 05-533, respectively) and anti-HA high-affinity antibody (Roche, Cat. no. 11867431001). R and separation magnet (Invitro10. Protein G DynaBeads gen, Cat. nos. 100.04D and 123.21D).

11. 7.5% SDS-PAGE gels. 12. Gel fixative solution (10% glacial acetic acid, 25% isopropanol, and 65% ddH2 O). 13. Amplify fluorographic reagent (GE Healthcare, Cat. no. NAMP100). 14. Gel dryer. 15. Phosphoscreen, phosphoimager, and image analysis software (ex. ImageJ. or ImageQuant).

3. Methods 3.1. TEPD Measurements in a Tracheal Xenograft Model

Newborn tracheas from CF and non-CF pigs and ferrets are obtained at birth sterilely and connected to flexible tubing through a series of adapters and stents to keep the trachea extended to normal length. These cassettes are then implanted subcutaneously into the flanks of athymic Nu/Nu mice (a host that will not reject the tissue). A schematic view of this ex vivo tracheal xenograft model is shown in Fig. 19.1. Importantly, these grafts become vascularized by 2–3 weeks and have ports that allow for lumenal access for TEPD measurements. Details on the methods for generating these cassettes and for the surgical implantation are described elsewhere (46). TEPD measurements of the xenografted tracheal airways can be used to assess changes in the permeability to various ions in response to antagonists and agonists of epithelial ion channels such as the epithelial sodium channel (ENaC) and CFTR. 1. After 4–5 weeks post-transplantation, the xenografts are fully differentiated and ready for TEPD analysis using the equipment shown in Fig. 19.2. The xenograft-bearing mouse (typically with two xenografts) is anesthetized by intraperitoneal injection of ketamine (100 mg/kg) and xylazine (20 mg/kg) in PBS. Once anesthetized, the mouse is placed on a sterile drape and the chrome wire caps from the xenograft exit ports are removed using sterile forceps. 2. The xenograft is gently flushed with 1 mL of Ham’s F12 medium using a 1-mL syringe and a butterfly needle.

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Fig. 19.1. Tracheal xenograft design and transplantation. (a) The cassette is composed of flexible plastic tubing, freshly excised newborn pig or ferret trachea, and chrome wire plugs (see Note 1). The trachea is fastened to the tubing using silk sutures. (b) The xenograft cassettes are inserted subcutaneously into the flanks of Nu/Nu athymic mice. The xenografts vascularize within 2–3 weeks and continue to mature and develop until ready for bioelectric characterization by measuring PD (4–5 weeks). PD recordings are made weekly until 8–9 weeks post-transplantation. Usually a CFTR–/– xenograft is transplanted in parallel to either a CFTR+/– or a CFTR+/+ xenograft.

Fig. 19.2. Potential difference instrumentation and setup. Measuring TEPD in this ex vivo model requires the following equipment: computer with data acquisition software, pH/mV meter, calomel electrodes, and syringe pump. The pH/mV is connected to the calomel electrodes that are connected to the anesthetized mouse by means of butterfly electrodes (see Note 2). The positive electrode is inserted into the perfusion tubing (black arrows) allowing access to the luminal surface of the trachea, while the negative electrode is inserted subcutaneously (white arrows).

3. The Ham’s F12 medium is removed by flushing the xenograft with air. This is done by removing the syringe from the butterfly needle, refilling it with air, returning the syringe to the butterfly needle, and gently forcing the air through the xenograft. 4. The syringe is removed and the butterfly needle is left on the distal port as a drain for the perfused TEPD buffer solutions.

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5. 10-mL syringes are filled with TEPD buffers and fitted on the syringe pump. A length of manifold pump tubing, which is to be placed on the needle of the first syringe, is first fitted onto the medial port of the xenograft by means of a shortened pipette tip (20–200 μL). 6. Calomel electrodes, immersed in 1 M KCl, are connected to the pH/mV meter and to the butterfly electrodes. 7. Butterfly electrodes are prepared by filling 21-gauge butterfly tubing with 5% noble agar in 1 M KCl (see Note 2). The positive butterfly electrode is inserted into the perfusion tubing just external to the xenograft port. This is done by directly inserting the 21-gauge needle through the tubing. The negative electrode is inserted subcutaneously in the back of the mouse. 8. Millivolt recordings are obtained from the pH/mV meter and data linked directly to a computer, equipped with data acquisition software. Measurements are taken every second. Example TEPD recordings and histological sections for ferret CFTR–/– and CFTR+/+ xenografts are depicted in Fig. 19.3. 9. Typically, the xenograft is sequentially perfused with 2 mL of each of the TEPD buffers through the syringe pump at a flow rate of 200 μL/min (10 min, see Notes 3 and 4). 10. After recording is complete, the xenograft is gently flushed with 1 mL of Ham’s F-12 medium using a butterfly needle and a 1-mL syringe. The syringe is then disconnected from the xenograft. The syringe is refilled with air by gently forcing air through the xenograft, restoring an air–liquid interface. The chrome wire inserts are replaced into the tubing ports. 11. Steps 1–10 are repeated (optional) on the xenograft on the other side of the animal if two xenografts are implanted. 12. Xenografts can be routinely measured up to two times per week and are typically irrigated with F12 media, followed by air, the day before each measurement to remove excess mucous. 3.2. Cross-Species Analysis of CFTR Processing Requires Antibodies that Efficiently Bind to Conserved Epitopes Between Species

When comparing biologic properties among different species of CFTR, it is imperative that the antibodies used react similarly across the species. This can be achieved by screening available antibodies against each species of CFTR to be compared and/or adding a common epitope tag such as HA to the fourth extracellular loop (which has not been shown to affect CFTR function). Summarized in Table 19.3 are results of an antibody screen for comparative studies between human and ferret CFTRs using Western blotting and immunoprecipitation. This section

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Fig. 19.3. TEPD analysis of ferret CF and non-CF tracheal xenografts. (a) Representative TEPD tracings of newborn ferret CFTR+/+ (dark line) and CFTR–/– (light line) tracheal xenografts. The buffer conditions change with ion channel agonists and antagonists are indicated above the tracing. (b) Reproducibility of sequential TEPD measurements taken in the same ferret CFTR+/+ and CFTR–/– xenografts at week intervals as indicated. Buffer conditions were the same as shown in (a) with the buffer number marked arrowheads. (c) Histological H&E sections of ferret CFTR+/+ (top) and CFTR–/– (bottom) xenografts. Note the intact pseudostratified ciliated epithelium (empty arrows) and the presence of submucosal glands (solid arrows) in both genotypes.

will focus on our methods of immunoprecipitation of CFTR using R and the subsequent detection by phosphorylation of DynaBeads CFTR with protein kinase A (PKA) and [γ–32 P]ATP as previously described with modifications (43, 48). A brief description of our Western blot protocol is contained in Note 5. 1. Immunoprecipitation a. Total protein (1 mg), from cell lysate derived from cells transiently transfected with human or ferret CFTR or EGFP expression plasmid (see Note 6), is aliquoted to a 2-mL Eppendorf tube and diluted to 1 mL using RIPA buffer containing freshly added protease inhibitors. Tween 20 is then added to a final concentration of 0.1%.

R&D Systems

CFC

Santa Cruz

24–1

Mr. Pink

H-182

N-Term

NBD1

C-Term

R Domain

Pre-NBD1

NBD1

NBD1

R Domain

NBD2

NBD1

R Domain

C-Term

N-Term

CFTR region

1:200

1:500

1:1000

1:1000

1:1000

1:500

1:1000

1:1000

1:1000

1:500

1:1000

1:1000

1:1000

WB dilution

+

+

+

+

++

++

++

+++

+++

+++

+++

+++

+++

WB human

+

+

+

+

++

–

++

+++

+++

+++

++

++

+++

WB ferret

+++

2 μg/mL

ND ND

7.5 μL ND

ND

ND ND

ND

ND +++

2 μg/mL

++

10 μg/mL ND

ND

ND

ND

ND

ND

ND

ND

+++

2 μg/mL ND

IP human

IP dilution

ND

ND

ND

ND

+++

ND

++

ND

ND

ND

ND

+++

+++

IP ferret

WB, Western blot; IP, immunoprecipitation; CFFT, cystic fibrosis foundation therapeutics; CFC, CFTR folding consortium; ND, not determined; –, no interaction; +, weak binding; ++, moderate binding; +++, strong binding. All WB and IPs came from a large pool of cell lysate derived from HT1080 cells expressing either human or ferret CFTR

R&D Systems

13–1

CFC

3G11

CFC

CFFT

570

Millipore

CFFT

596

L12B4

CFFT

660

10B6.2

Millipore

CFFT

MM13-4

Millipore

M3A7

217

Source

Name

Table 19.3 Human and ferret CFTR antibody optimization

326 Fisher, Zhang, and Engelhardt

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b. Primary antibody is incubated with the dilution specified in Table 19.3 for at least 2 h at 4◦ C with rotation. R (50 μL) is added to the Washed protein G DynaBeads solution and the mixture is incubated overnight at 4◦ C (see Notes 7 and 8). c. The beads are washed three times with PBS containing 0.1% Tween 20, placing the tube on a magnet between each wash to remove the supernatant. 2. In vitro phosphorylation of CFTR: a. Prepare PKA and ATP phosphorylation buffers. b. The beads from the finished IP protocol are washed once with PBS and the supernatant is removed by placing the tube on the magnet. c. The beads are resuspended in 20 μL of the PKA phosphorylation buffer. d. Add 40 μL of the ATP phosphorylation buffer and shake for 30 min at 30◦ C (see Note 9). e. The beads are washed four times with PBS containing 0.1% Tween 20. f. Beads, antibody, and CFTR are dissociated by adding 2× SDS-PAGE loading buffer, incubating at 37◦ C with continual shaking. g. Place the tube in the magnet and load the supernatant on a 7.5% SDS-PAGE gel. h. Resolve by electrophoresis (see Note 10). i. Fix the gel by transferring the gel to a disposable plastic container filled with fixative solution (10% glacial acidic acid, 25% isopropanol, and 65% ddH2 O) and gently agitate for 30 min. j. Dry the gel by transferring to a piece of filter paper, cover with plastic wrap, and dry on a vacuum gel drier at 80◦ C for 45 min. k. Expose to a pre-cleared phosphoscreen and scan several days later using a phosphoimager. 3.3. Cross-Species Analysis of the Processing Efficiency and Stability of CFTR by [35 S]Methionine and Cysteine Pulse Chase

The glycosylation characteristics of CFTR serve as an excellent endpoint for assessing the processing CFTR. The immature form of CFTR resides in the endoplasmic reticulum (ER) and is called band B (∼150 kDa), while the fully glycosylated mature protein resides at the plasma membrane and is called band C (∼170– 180 kDa). Note that the migratory apparent molecular weight of CFTR can vary slightly across species and may be due to slightly altered glycosylation. Metabolic pulse-chase experiments using radioactive amino acids, [35 S]methionine and [35 S]cysteine,

328


Fig. 19.4. Metabolic [35 S]methionine pulse chase of ferret CFTR processing. (a) HT1080 cells transiently expressing ferret CFTR were starved of methionine and cysteine (30 min), labeled with [35 S]methionine and [35 S]cysteine (15 min), and chased with media containing cold methionine and cysteine for the given time points as described under Section 3.3. (b) Densitometric quantification of (a) Empty points representing band BT /band B0 × 100. Solid data points representing band CT /band B0 × 100 (see Note 13).

serve as an excellent method to characterize the rate, stability, and efficiency of processing from band B to C of wild-type and mutant CFTRs across species. An example of a metabolic pulsechase autoradiograph and quantification thereof for ferret wildtype CFTR is shown in Fig. 19.4. 1. Depletion of intracellular pools of methionine and cysteine: Gently wash the cells expressing CFTR two times with warm starvation media. Incubate the cells at 37◦ C for 30 min in starvation media (2 mL). 2. Metabolic labeling of CFTR: Gently aspirate the starvation media. Transfer cells to an area designated for the use of radioactive materials (see Note 9). Add 2 mL of starvation media containing 0.2 mCi/mL of [35 S]methionine and cysteine. Incubate at 37◦ C for 15 min. 3. Metabolic chase of labeled CFTR: Aspirate the labeling media and place the cells on ice. Wash three times in cold PBS. Add warm chase media (4 mL) and incubate at 37◦ C for the desired amounts of time (see Note 11). 4. Harvesting the cells: Place cells on ice and remove the chase media by aspiration. Proceed to gently wash the cells three times with ice-cold PBS (1 mL). Lyse the cells on ice in 1 mL of RIPA buffer containing protease inhibitors for 5–30 min. Transfer the lysate to a 1.5-mL Eppendorf tubes and spin at 16,000×g for 10 min at 4◦ C (see Note 12). Freeze the samples until last chase time point has been harvested.

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5. Immunoprecipitation of labeled CFTR: Thaw lysate and add Tween 20 to a final concentration of 0.1%. Then add 2 μg of anti-CFTR antibodies M3A7, L12B4, and MM13-4 and rotate for 3–5 h at 4◦ C. Wash 50 μL (30 μg/mL) of R with PBS containing 0.1% Tween 20, pelletDynaBeads ing between each wash. Resuspend the beads in the original volume and add to the antibody/antigen containing lysate. Rotate at 4◦ C overnight. Wash the lysate with PBS (0.1% Tween 20) six times. Dissociate the protein and antibody by adding 2× SDS loading buffer (30 μL) and shake at 37◦ C for 30 min. 6. Electrophoresis and autoradiography: Load the lysate on 7.5% SDS-PAGE gels and electrophorese overnight (30 V). Fix the gel for 30 min by gently shaking at room temperature in the fixative solution. Replace the fixative solution with Amplify and incubate for 30 min at room temperature with gentle shaking. Dry the gel using a gel dryer and expose to a phosphoscreen for several days. Develop/scan the phosphoscreen using a phosphoimager and imaging software. 7. Densitometric analysis: Quantify the total CFTR signal for bands B and C for each of the experimental time points (see Note 13).

4. Notes 1. Freshly isolated trachea from newborn ferret or pig is cannulated to the previously described xenograft cassette (46). In brief, the cassette consists of a combination of silastic (Dow Corning, Midland, MI) and Teflon (Thomas Scientific, Swedesboro, NJ) tubing attached to barb-to-barb connectors (Bio-Rad Laboratories, Hercules, CA). The trachea is ligated to the connectors and the tubing ports capped with chromel A steel wire (Hoskins MFG, Novi, MI). These cassettes are inserted subcutaneously into the flanks of male Nu/Nu athymic mice. 2. Butterfly electrodes are made by dissolving the agar noble (5%) in 1 M KCl by heating. The butterfly needle/tubing is filled with this hot solution by negative pressure generated by a 30-mL syringe. These electrodes are submerged in 1 M KCl in a 100-mm dish for up to 1 year at 4◦ C. 3. To accurately calculate changes in potential difference between buffers, it is important to wait for a stable millivolt reading before switching to the next buffer. This usually occurs within 10 min but depends upon the xenograft.

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4. It is critical that the fluid-filled tubing and the xenograft are devoid of any air bubbles. The manifold tubing is therefore clamped with a hemostat when changing buffers. The hemostat is removed once the tubing is attached to the next buffer. Failure to keep air from the system will result in moments of infinite TEPD spikes as the air moving through the xenograft disrupts the electrical conductivity. 5. Equal amounts of protein from cellular lysate from cells expressing human or ferret CFTR (see Note 6) were resolved electrophoretically on 6% SDS-PAGE gels. The protein was transferred to nitrocellulose membranes and the membrane blocked in PBS containing 0.1% casein and 0.2% Tween 20. CFTR primary antibodies were then added at the dilution indicated in Table 19.3 and incubated for 1 h at room temperature. The blots were washed, probed with secondary antibodies conjugated to an IR dye, and imaged using an Odyssey IR scanner. 6. Transient expression of human and ferret CFTRs in HT1080 cells was achieved by electroporation using the BTX-830/630B system (Harvard Apparatus, Holliston, MA). The following electroporation conditions were used for HT1080 (4 pulses, 230 mV, 1 ms interval) and BHK21 (1 pulse, 260 mV, 1 ms interval) cell lines. R 7. Incubation of the antigen/antibody and the DynaBeads for 2 h at room temperature is also sufficient. It is also critR be maintained in a 0.01–0.1% ical that the DynaBeads Tween 20 solution to avoid bead clumping.

8. Alternatively, one can prebind the CFTR antibodies and R by incubating with rotation for 30 min at the DynaBeads room temperature. The unbound antibody is then washed R can be directly away and the antibody-bound DynaBeads added to the cell lysate and rotated for 2 h at room temperature. 9. From this step forward, everything must be carried out in the radiation room according to manufacturer’s guidelines. Everything must be discarded appropriately and appropriate personal protective equipment used. 10. To achieve adequate separation between bands B and C on a 7.5% mini gel, we run the ladder off to 100 kDa. This is done by running the gel at 120 V for ∼3 h or 30 V overnight. 11. It is recommended that several time points from 0 to 12 h be included to adequately study the rate of disappearance of band B and appearance and disappearance of band C. It may be helpful to also include time points up to 48 h to ascertain differences in protein stability.

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12. To avoid non-specific pull down of other labeled proteins, keep the lysis time less than 30 min and spin the lysate at 4◦ C. This avoids protein degradation and lysis of nuclei in the sample. 13. Quantify the lane background using two regions for each lane (below band B and above band C). Subtract the lane background from the regions representing bands B and C. To analyze the disappearance of band B, measure the percentage of band B over time divided by the initial amount of band B (band BT /band B0 ). To analyze the appearance/disappearance of the higher molecular weight, fully processed band C, it is important to measure the intensity of band C over time divided by the initial band B labeling intensity (band CT /band B0 ). Due to background issues throughout the lane at time 0 of the chase (likely due to undegraded fragments of CFTR generated during the labeling process), band C0 is set to 0.

Acknowledgments This work was supported by grants from the NHLBI (RC1HL099516), NIDDK (P30DK054759, R37DK047967), NHLBI (HL091842) and the Cystic Fibrosis Foundation (ENGELH08XX0), as well as by the Roy J. Carver Chair in molecular medicine. We also gratefully acknowledge Drs Christine Blaumueller and Monali Sawai for editorial contributions. References 1. Davidson, D. J., and Dorin, J. R. (2001) The CF mouse: an important tool for studying cystic fibrosis. Expert Rev Mol Med 3, 1–27. 2. Davidson, D. J., and Rolfe, M. (2001) Mouse models of cystic fibrosis. Trends Genet 17, S29–S37. 3. Dickinson, P., Dorin, J. R., and Porteous, D. J. (1995) Modelling cystic fibrosis in the mouse. Mol Med Today 1, 140–148. 4. Egan, M. E. (2009) How useful are cystic fibrosis mouse models? Drug Discovery Today: Disease Models 6, 35–41. 5. Grubb, B. R., and Boucher, R. C. (1999) Pathophysiology of gene-targeted mouse models for cystic fibrosis. Physiol Rev 79, S193–S214. 6. Sun, X., Yan, Z., Yi, Y., Li, Z., Lei, D., Rogers, C. S., et al. (2008) Adeno-associated

virus-targeted disruption of the CFTR gene in cloned ferrets. J Clin Invest 118, 1578–1583. 7. Rogers, C. S., Hao, Y., Rokhlina, T., Samuel, M., Stoltz, D. A., Li, Y., et al. (2008) Production of CFTR-null and CFTR-DeltaF508 heterozygous pigs by adeno-associated virus-mediated gene targeting and somatic cell nuclear transfer. J Clin Invest 118, 1571–1577. 8. Dorin, J. R., Dickinson, P., Alton, E. W., Smith, S. N., Geddes, D. M., Stevenson, B. J., et al. (1992) Cystic fibrosis in the mouse by targeted insertional mutagenesis. Nature 359, 211–215. 9. Snouwaert, J. N., Brigman, K. K., Latour, A. M., Malouf, N. N., Boucher, R. C., Smithies, O., et al. (1992) An animal model for cystic

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10.

11.

12.

13.

14.

15.

16.

17.

18.

19.

20.

Fisher, Zhang, and Engelhardt fibrosis made by gene targeting. Science 257, 1083–1088. O’Neal, W. K., Hasty, P., McCray, P. B., Jr., Casey, B., Rivera-Perez, J., Welsh, M. J., et al. (1993) A severe phenotype in mice with a duplication of exon 3 in the cystic fibrosis locus. Hum Mol Genet 2, 1561–1569. Ratcliff, R., Evans, M. J., Cuthbert, A. W., MacVinish, L. J., Foster, D., Anderson, J. R., et al. (1993) Production of a severe cystic fibrosis mutation in mice by gene targeting. Nat Genet 4, 35–41. Zhou, L., Dey, C. R., Wert, S. E., DuVall, M. D., Frizzell, R. A., and Whitsett, J. A. (1994) Correction of lethal intestinal defect in a mouse model of cystic fibrosis by human CFTR. Science 266, 1705–1708. Colledge, W. H., Abella, B. S., Southern, K. W., Ratcliff, R., Jiang, C., Cheng, S. H., et al. (1995) Generation and characterization of a delta F508 cystic fibrosis mouse model. Nat Genet 10, 445–452. Hasty, P., O‘Neal, W. K., Liu, K. Q., Morris, A. P., Bebok, Z., Shumyatsky, G. B., et al. (1995) Severe phenotype in mice with termination mutation in exon 2 of cystic fibrosis gene. Somat Cell Mol Genet 21, 177–187. van Doorninck, J. H., French, P. J., Verbeek, E., Peters, R. H., Morreau, H., Bijman, J., et al. (1995) A mouse model for the cystic fibrosis delta F508 mutation. EMBO J 14, 4403–4411. Zeiher, B. G., Eichwald, E., Zabner, J., Smith, J. J., Puga, A. P., McCray, P. B., Jr., et al. (1995) A mouse model for the delta F508 allele of cystic fibrosis. J Clin Invest 96, 2051–2064. Delaney, S. J., Alton, E. W., Smith, S. N., Lunn, D. P., Farley, R., Lovelock, P. K., et al. (1996) Cystic fibrosis mice carrying the missense mutation G551D replicate human genotype–phenotype correlations. EMBO J 15, 955–963. Rozmahel, R., Wilschanski, M., Matin, A., Plyte, S., Oliver, M., Auerbach, W., et al. (1996) Modulation of disease severity in cystic fibrosis transmembrane conductance regulator deficient mice by a secondary genetic factor. Nat Genet 12, 280–287. Dickinson, P., et al. (1998) Generation of a CF mutant mouse possessing the G480C mutation. In 22nd European CF Conference Berlin Book of Abstracts, PS7. 14, 143. Du, M., Liu, X., Welch, E. M., Hirawat, S., Peltz, S. W., and Bedwell, D. M. (2008) PTC124 is an orally bioavailable compound that promotes suppression of the human

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Fisher, Zhang, and Engelhardt atresia in fetuses and neonates. Pediatr Pathol Lab Med 16, 25–40. Oppenheimer, E. H., and Esterly, J. R. (1962) Observations in cystic fibrosis of the pancreas. II. Neonatal intestinal obstruction. Bull Johns Hopkins Hosp 111, 1–13. Bronstein, M. N., Sokol, R. J., Abman, S. H., Chatfield, B. A., Hammond, K. B., Hambidge, K. M., et al. (1992) Pancreatic insufficiency, growth, and nutrition in infants identified by newborn screening as having cystic fibrosis. J Pediatr 120, 533–540. Weber, A. M., Roy, C. C., Chartrand, L., Lepage, G., Dufour, O. L., Morin, C. L., et al. (1976) Relationship between bile acid malabsorption and pancreatic insufficiency in cystic fibrosis. Gut 17, 295–299. O’Brien, S., Mulcahy, H., Fenlon, H., O’Broin, A., Casey, M., Burke, A., et al. (1993) Intestinal bile acid malabsorption in cystic fibrosis. Gut 34, 1137–1141. Lindblad, A., Glaumann, H., and Strandvik, B. (1999) Natural history of liver disease in cystic fibrosis. Hepatology 30, 1151–1158. Feranchak, A. P., and Sokol, R. J. (2001) Cholangiocyte biology and cystic fibrosis liver disease. Semin Liver Dis 21, 471–488. Chaudry, G., Navarro, O. M., Levine, D. S., and Oudjhane, K. (2006) Abdominal mani-

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Chapter 20 CFTR Folding Consortium: Methods Available for Studies of CFTR Folding and Correction Kathryn W. Peters, Tsukasa Okiyoneda, William E. Balch, Ineke Braakman, Jeffrey L. Brodsky, William B. Guggino, Christopher M. Penland, Harvey B. Pollard, Eric J. Sorscher, William R. Skach, Philip J. Thomas, Gergely L. Lukacs, and Raymond A. Frizzell Abstract The CFTR Folding Consortium (CFC) was formed in 2004 under the auspices of the Cystic Fibrosis Foundation and its drug discovery and development affiliate, CFF Therapeutics. A primary goal of the CFC is the development and distribution of reagents and assay methods designed to better understand the mechanistic basis of mutant CFTR misfolding and to identify targets whose manipulation may correct CFTR folding defects. As such, reagents available from the CFC primarily target wild-type CFTR NBD1 and its common variant, F508del, and they include antibodies, cell lines, constructs, and proteins. These reagents are summarized here, and two protocols are described for the detection of cell surface CFTR: (a) an assay of the density of expressed HA-tagged CFTR by ELISA and (b) the generation and use of an antibody to CFTR’s first extracellular loop for the detection of endogenous CFTR. Finally, we highlight a systematic collection of assays, the CFC Roadmap, which is being used to assess the cellular locus and mechanism of mutant CFTR correction. The Roadmap queries CFTR structure– function relations at levels ranging from purified protein to well-differentiated human airway primary cultures. Key words: Protein folding, protein degradation, antibody generation, cell surface protein detection, research consortium, www.cftrfolding.org.

K.W. Peters and T. Okiyoneda contributed equally to this chapter


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1. Introduction 1.1. The CFTR Folding Consortium (CFC): Rationale

CFTR and many of its disease mutants are prominent substrates for endoplasmic reticulum-associated degradation (ERAD). As a multi-subunit ABC transporter protein, the biogenesis of CFTR is a complex process that involves multiple positive (pro-folding) and negative (pro-degradation) protein interactions. The common disease mutation (F508del), which results in the omission of phenylalanine at position 508 (a class II mutation), is characterized by defective biogenesis and near-complete ERAD (1). Approximately 30,000 CF patients reside in North America and more than 90% carry the F508del mutation on at least one allele; ~50% are F508del homozygotes. The F508del mutation produces severe disease, with a life expectancy of only 24 years, as opposed to 37 years in the general patient population (CF Fdn 2005 Patient Registry). Thus, among the many mutations responsible for CF, correction of the molecular defect imposed by F508del offers the maximal potential for improving the quality of life and life expectancy of CF patients. Interactions of CFTR with folding/degradation pathway components are determined by the protein’s conformation, which can be monitored by comparing the proteolytic cleavage patterns of the WT and mutant proteins. Such studies have indicated that the protease cleavage patterns are similar for immature WT and F508del proteins, whereas the digestion pattern of mature WT-CFTR is more compact, reflecting its folded state (2, 3). These data support the concept that ER-retained F508del-CFTR achieves intermediate conformation(s) that lie along the normal CFTR-folding pathway, and they suggest that misfolding arrests F508del at one or more critical checkpoints. This concept implies that F508del can be rescued from ERAD if the limiting step(s) are appropriately manipulated. Indeed, F508del-CFTR can be rescued biochemically and functionally by low temperature (4), chemical chaperones (5), chaperone manipulations (6–8), intragenic suppressor mutations (9, 10), or ER retention motif mutations (11). On this basis, and beginning approximately 10 years ago, the Cystic Fibrosis Foundation linked with biotechnology firms and academic researchers to utilize high-throughput screening (HTS) as a drug discovery platform to identify small molecules that promote F508del-CFTR trafficking to the plasma membrane (12, 13). By definition, small molecule correctors would facilitate the delivery of functional CFTR to the surface of epithelial cells. Nevertheless, HTS platforms provide end-point assays of CFTR function (e.g., anion efflux or membrane voltage) or of cell surface protein expression, which do not provide information on the

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mechanism by which additional CFTR has progressed to the cell surface. Given the complex series of interactions that may rescue mutant CFTR, knowledge of the molecular target(s) may be important in the advancement of drugs through approval processes, in the evaluation of potential off-target effects (i.e., toxicity), and in the potential development of combination therapies that would target different pathways. In addition, a byproduct of the drug discovery process is often the provision of new tools for a vertical evaluation of mechanism of action and for judging selectivity within a protein class. Compounds on the critical path for human therapy that do not define a mechanism of action do not address these important issues. Accordingly, it became apparent that furthering our understanding of the mechanisms whereby misfolded proteins can be progressed through the secretory pathway would benefit from coordinating the efforts of multiple investigators with complementary assays and areas of expertise that report on distinct aspects of CFTR biology. This CFTR Folding Consortium has now expanded to 10 academic laboratories, comprising the authorship of this chapter. Key goals of CFC include (a) the generation of new tools and assays for investigating CFTR folding and biogenesis and the means to share them with the CF scientific community and (b) the identification of cellular pathways that mediate CFTR processing as targets for the potential therapeutic manipulation of F508del-CFTR mis-processing. Accordingly, the unifying hypothesis linking these goals states that the unproductive course of F508del-CFTR biogenesis can be overcome by understanding and manipulating the intra-molecular fold and/or the rate-limiting inter-molecular interactions required for F508del trafficking to, and function at, the cell surface. 1.2. CFC Web Site

The methods and reagents developed by the CFC are made available to researchers focused on the above hypothesis through the consortium Web site: www.cftrfolding.org. In the spirit of the consortium effort, investigators who utilize these resources are asked to provide feedback on their utility and to make available any new and improved reagents or methods developed from them. Reagents available at present include antibodies (primarily to NBD1), NBD1 proteins, cell lines, and expression vectors for related proteins and shRNAs. Protocols for the use of these reagents are available from CFC investigators, and their posting on the Web site is an ongoing process.

1.3. CFTR Reagents

While there are many useful antibodies available for CFTR detection and localization, our understanding of CFTR folding would benefit from the availability of antibodies whose interaction with the protein is conformation dependent, permitting them to distinguish between wild-type CFTR and its F508del variant. Several

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CFC laboratories have been actively pursuing this goal by raising antibodies to NBD1. Currently, five NBD1 antibodies are available through the Web site, including the mouse monoclonals, 5A6.3, 10B6.2, and 7D12; the rat monoclonal, 3G11; and the rabbit polyclonal, Mr. Pink. Interestingly, the majority of the monoclonal antibodies recognize a common epitope, which lies within aa 401–410 of hNBD1, while the available polyclonal targets several NBD1 epitopes, as would be expected. Further attempts to generate antibodies sensitive to conformation continue, using CFTR domains or full-length protein as antigens. Success in this endeavor may be reflected in antibodies that recognize the folded protein in immunoprecipitation or immunofluorescence experiments but detect the unfolded protein less well in Western blots or vice versa. Such tools would permit the evaluation of conditions and agents that improve the folding of mutant CFTR. 1.3.2. CFTR Proteins

Purified mouse and human NBD1 proteins are available for distribution to investigators. Wild-type and F508del-mNBD1 and WT-hNBD1 (residues 389–673) are provided for shipping costs in aliquots of a few hundred micrograms. Requests for larger amounts can be accommodated with justification. Requests for the reagents can be placed at the Folding Consortium Web site under “Reagents.” A complete set of characterization data for each preparation (SDS-PAGE, CD spectrum, intrinsic tryptophan fluorescence spectrum, and thermal stability, as described in Chapter 20) is also included on the Web site under the “Data” tab. Identity of the expressed protein has been verified by mass spectrometry for samples of each of the preparations as well. Protocols used for the expression, purification, and characterization are also available on the Web site under the “Protocols” tab. Whereas the F508del-hNBD1 is more difficult to produce, due to decreased stability, it is not routinely available via the CFC Web site.

1.3.3. The CFC Roadmap

A goal of the Folding Consortium is to combine the diverse array of cell-based and in vitro assays available in member labs to probe diverse aspects of CFTR biogenesis and trafficking. To systematize this process, the collection of assays was overlaid on a modified scheme for the protein secretory pathway, featuring intersections that allow distinct aspects of CFTR function to be assessed. Thus, the Roadmap is basically an assay-laden decision tree that reflects the cellular fate of CFTR. The analysis of CFTR function at each of these nodes provides a fingerprint of how a specific modulator (potentiator or corrector) influences CFTR’s function at critical sites of the secretory pathway. For example, macroscopic assays of transepithelial currents across airway epithelia provide


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an indication of whether pre-incubation with a compound rescues F508del-CFTR function in the context of a relevant cellular background. Assays performed at other nodes, approximately 30 in all, determine whether this action can be attributed to increased cell surface protein expression, improved intracellular trafficking, reduced ERAD, or the folding of CFTR toward a native conformation. Depending on the outcome, in-depth assays are available to assess the pathways that are responsible for the observed effect(s). Details of the Roadmap and its attendant assays are provided in a manuscript that focuses on the analysis of CFTR modulators (14). That report summarizes the potential uses of Roadmap assays and provides a comparative analysis of the actions of available small molecule correctors, reduced temperature, and revertant mutations that improve the F508del-CFTR fold. The Modulator Roadmap provides an assay-based view of the protein secretory pathway, rather than the one found in textbooks. Due to the diversity of its assays, this analysis provides a more complete picture of mechanism than can generally be obtained by individual laboratories or biotechnology concerns, at least within a timely manner. Nevertheless, each of the map intersections is determined by pathways involving hundreds of proteins and protein interactions, requiring a more complete systems biology analysis of the actions of the most efficacious compounds. A key implication from the findings to date is that different F508del correction approaches show activity profiles that allow their distinction from one another (14). Potentially, this outcome could inform drug development by (a) identifying ratedetermining steps in the complex process of CFTR biogenesis; (b) informing structure–activity relationships by associating different drug scaffolds with activity at the same or different nodes within the Roadmap; (c) implicating compound interactions that may lead to additivity or synergy when treatments are combined; and (d) suggesting candidates for the molecular target(s) of drug action. In general, the profile of drug activity in this analysis may provide for comparisons of different CFTR correction strategies to optimize our efforts to understand and treat the most common cause of cystic fibrosis. In addition, this approach may be generally applicable to other diseases of protein folding. 1.4. Methods to Determine CFTR Membrane Expression 1.4.1. Measuring CFTR Cell Surface Density by Cell Surface ELISA

The cell surface density of CFTR can be determined by biotinylation combined with immunoprecipitation and immunoblotting (15–17). However, this experimental approach is technically demanding and time consuming and has a relatively low sensitivity. Exogenous and endogenous CFTR could also be detected at the plasma membrane by antibodies recognizing extracellular segments of the channel (e.g., the first extracellular loop)

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(18, 19). We have developed a third approach using cell surface ELISA (designated as immunoperoxidase assay) to monitor quantitatively the cell surface expression level, as well as turnover of heterologously expressed wild-type and mutant CFTR bearing an extracellular epitope tag (20, 21) (Fig. 20.1a). To this end, a 3HA epitope tag was introduced genetically into the fourth extracellular loop of CFTR (CFTR-3HA) (12, 20). CFTR variants with the 3HA tag were expressed heterologously in BHK, HEK293, HeLa, IB3, and CFBE cells. We confirmed that the

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Fig. 20.1. (a) Schematic model of the immunoperoxidase assay to measure CFTR cell surface density, internalization rate, and cell surface stability. The CFTR cell surface density is measured by the primary anti-HA and secondary HRP-conjugated antibody binding to the CFTR-3HA. The HRP activity is monitored by the fluorescence generated R Red substrate. CFTR cell surface stability and internalization rate from the Amplex are quantified by measuring the disappearance of the anti-HA antibody from the cell surface at 37◦ C following the primary antibody binding. The remaining anti-HA antibody is quantified by the immunoperoxidase assay. (b) Cell surface density of CFTR-3HA in HEK293MSR cells. Similar to previous studies (12, 20, 21), most F508del-CFTR failed to express at the cell surface. Low-temperature incubation (26◦ C, 36 h) rescued the cell surface expression of F508del-CFTR (rF). (c) Internalization rate (5 min) of the wt and rescued F508del-CFTR-3HA in HEK293MSR cells. (d) Cell surface stability of CFTR-3HA in HEK293MSR cells was measured by the immunoperoxidase assay. While the wt-CFTR was stable at the cell surface, rF508del-CFTR was rapidly internalized (c) and eliminated from the cell surface (d).


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3HA tag has minimal effect on CFTR folding, channel functioning, and trafficking (12, 20–22). The wild-type (wt) and mutant CFTR-3HA expression, as well as internalization and stability at the plasma membrane (PM), was detected by the immunoperoxidase assay using primary antiHA antibody and HRP-conjugated secondary antibody in the R Red (Invitrogen), a florescent HRP subpresence of Amplex strate. Normalization of the fluorescence signal for cell number or cellular protein permits the quantitative comparison of the channel density under various conditions, as well as the PM turnover rate of CFTR, an assay described in Section 3.1.1. CFTR internalization and cell surface stability are followed by a modified version of the immunoperoxidase assay as described in Section 3.1.2. 1.4.2. Cell Surface Detection of Native CFTR

Epitope or fluorescent protein tagging of CFTR has been remarkably successful, permitting the cell surface labeling of expressed protein for a variety of purposes, including the screening for small molecule correctors of mutant CFTR trafficking (23, 24). Nevertheless, the detection of native, untagged CFTR at the plasma or the apical membrane remains a significant goal, particularly in epithelia that express the channel endogenously. If successful, this approach would have the benefit of eliminating over-expression artifacts and cell-type dependency in the cellular handling of WT and mutant CFTR. High-affinity antibodies that recognize an extracellular epitope of endogenous CFTR would be particularly useful in preclinical studies to evaluate potential therapies to promote CFTR progression to the apical plasma membrane. However, only 4% of CFTR’s amino acids are predicted to be located at the extracellular surface, and the majority of these are likely to be shielded by glycosylation at two sites in the larger fourth extracellular loop. However, the first ECL contains 15 amino acids in a sequence predicted to be antigenic. Amino acids 107–118 were used previously to raise antibodies, and they detected CFTR in the plasma membrane of non-permeabilized cells, with amplification procedures (18, 19). They provided some of the first evidence that F508del-CFTR did not appear at the cell surface. This antibody was less efficient in immunoprecipitation experiments, perhaps because it did not detect the native conformation of ECL1 at high affinity. In anticipation of raising higher affinity antisera, we employed a conformationally constrained ECL1 peptide as antigen, in which the N and C termini were linked by a disulfide bond. Results obtained with this rabbit antibody in Western blot, immunoprecipitation, and immunofluorescence studies are described in Section 3.2.

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2. Materials 2.1. Materials for Detection of HA-CFTR

1. Cell culture medium: Use the appropriate bicarbonatecontaining medium (e.g.„ Dulbecco’s modified Eagle’s medium (DMEM) for HeLa or HEK293 cells, DMEM/F12 for BHK cells) supplemented with 10% FBS (Invitrogen) for culturing the cells in a CO2 incubator. Medium should be stored at 4◦ C. 2. PBS(+): Phosphate-buffered saline supplemented with 0.1 mM CaCl2 and 1 mM MgCl2 . 3. 0.5% Bovine serum albumin (BSA)/PBS(+). Store at 4◦ C. 4. Anti-HA11 monoclonal antibody (Covance MMS-101R). 5. HRP-conjugated anti-mouse IgG secondary antibody (GE Healthcare NA931V). R Red reagent (Invitrogen A-22177, 10 mM 6. Amplex DMSO stock, stored at –20◦ C, protect from light).

7. 30% H2 O2 (Sigma H1009). R R Red reaction mix (50 μM Amplex Red, 8. Amplex 200 μM H2 O2, 50 mM NaH2 PO4 , pH 7.4, protect from light, prepare immediately before use).

9. RIPA buffer (150 mM NaCl, 20 mM Tris, 1% Triton X100, 0.1% SDS, 0.5% sodium deoxycholate, pH 8.0). 10. BCA protein assay kit (Thermo Scientific, #23225). 2.1.1. Instrumentation

1. Tissue culture incubator at 37◦ C with 5% CO2 . 2. 24-Well tissue culture plates (BD Falcon DL-353047). 3. 96-Well black plates for fluorescence (NUNC 437111). 4. Fluorescence plate reader (POLAR star Optima, BMG LABTECH).

2.2. Materials for Detection of Native CFTR

1. Cell culture and lysis: Wild-type cystic fibrosis bronchial epithelial (wt-CFBE) (a kind gift of J.P. Clancy) cells are cultured in growth medium consisting of minimum essential medium (MEM) (Invitrogen) supplemented with 10% fetal bovine serum (FBS) (HyClone), 50 U/ml penicillin, 50 μg/ml streptomycin, 2 mM L-glutamine (Invitrogen), and 0.5 μg/ml puromycin (InvivoGen). F508delCFBE cells are cultured as wt-CFBEs but with 2 μg/ml puromycin. Parental CFBE cells (p-CFBEs) are cultured as wt-CFBEs but without puromycin. Calu-3 cells are cultured in MEM (ATCC), 15% FBS (HyClone), and 1% penicillin/streptomycin (Gibco). A 0.25% trypsin–0.53 mM EDTA solution (ATCC) is used to remove cells from plastic.


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2. 4-(2-Hydroxyethyl)-1-piperazine ethanesulfonic acid (HEPES)-buffered saline, pH 7.4–7.6 (HBS): 19 mM HEPES, 122 mM NaCl, 1 mM glucose, 6.3 mM Na2 HPO4 •7H2 O (dibasic), and 250 μl 0.5% phenol red solution. 10% Polyvinylpyrrolidone (PVP) (Sigma) is made in HBS. 3. 0.2% Ethylene glycol-bis(2-aminoethyl-ether)-N,N,N ,N tetraacetic acid (EGTA) is made in HBS. 4. Fifty milliliter of PVP/EGTA/trypsin (PET) is made with 35 ml HBS, 5 ml of 10% PVP, 5 ml of 0.2% EGTA, and 5 ml of 0.25% trypsin with Versene (Sigma). 5. Lysis buffer (RIPA): 150 mM NaCl, 50 mM Tris–HCl (pH 7.5), 1.0% Triton X-100, 1% deoxycholic acid (sodium salt), 0.1% sodium dodecyl sulfate (SDS), Complete Mini protease inhibitor tablet (PIT; Roche) (one tablet/10 ml RIPA). 2.2.1. Coupling of Peptide

1. Cognate sequence of the first extracellular loop of CFTR with three additional cysteines. R maleimide-activated mariculture keyhole limpet 2. Imject hemocyanin (mcKLH) (Pierce). R (Pierce) dialysis cassettes. 3. Slide-A-Lyzer

2.2.2. Production of Polyclonal Antibodies

1. Synthetic peptide coupled with KLH. 2. PBS (Gibco). R 3. Imject Freund’s complete adjuvant (Pierce). R Freund’s incomplete adjuvant (Pierce). 4. Imject

5. Two or three 6-week-old male rabbits. 2.2.3. Enzyme-Linked Immunosorbent Assay (ELISA)

1. Carbonate buffer: 15 mM Na2 CO3 , 35 mM NaHCO3 , 3 mM NaN3 , pH 9.6 (25). 2. Diethanolamine buffer: 1 M diethanolamine, 3 mM NaH3 , 0.5 mM MgCl2 , pH 9.8, stored in the dark. 3. Tris buffered saline (TBS): 50 mM Tris–HCl, 0.2 M NaCl, pH 7.5. 4. TBS/Tween (TBST): 50 mM Tris–HCl, 0.2 M NaCl, pH 7.5, 0.05% Tween 20. 5. TBST/bovine serum albumin (TBSTB): 50 mM Tris–HCl, 0.2 M NaCl, pH 7.5, 0.05% Tween 20, 1.0% bovine serum albumin. 6. Rabbit serum expressing antibodies of interest. 7. Alkaline phosphatase-conjugated AffiniPure goat anti-rabbit IgG (Jackson ImmunoResearch).

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2.2.4. Immunoprecipitation (IP)

1. Protein A agarose (Invitrogen) (26).

2.2.5. Immunoblot (IB)

1. Tris buffered saline (TBS): 50 mM Tris–HCl, 0.2 M NaCl, pH 7.5.

2. Sample buffer: 62.5 mM Tris–HCl (pH 6.8), 2% SDS, 10% glycerol, 0.01% bromophenol blue, and 5% β-mercaptoethanol.

2. TBS/0.1% Tween 20 (TBST). 3. TBST/5% instant non-fat dry milk (Carnation) (TBSTM). 4. Chemiluminescent reagents (GE Healthcare). 2.2.6. Immunofluorescence (IF)

1. Calu-3 cells on filters. 2. PBS. 3. Agonist: 10 ml PBS with 10 μl of 10 mM forskolin (Calbiochem) (fsk) in EtOH and 20 μl of 500 mM 3-isobutyl1-methylxanthine (IBMX) (Sigma) in dimethyl sulfoxide (DMSO). 4. PBS/0.5% bovine serum albumin (BSA)/0.15% glycine (PBSBG). 5. PBSBG/5% goat serum (PBSBG-GS).

3. Methods 3.1. Methods for Detection of Cell Surface HA-CFTR 3.1.1. Cell Surface Density Measurement of CFTR-3HA

1. Seed cells expressing CFTR-3HA on 24-well plates at least 24 h before the experiment. Prepare three or four wells of each sample so that triplicate or quadruplicate measurements could be obtained. Prepare non-transfected cells to verify the extent of the non-specific binding of the primary and the secondary antibody. By the time of the experiment cells should be at 80–90% confluence. 2. Rinse cells with ice-cold 0.5% BSA–PBS(+) (1 ml/well) gently to avoid cell loss. 3. Add ice-cold 0.5% BSA–PBS(+) (1 ml/well) and block in the same medium for 20 min on ice. 4. Bind the primary anti-HA antibody (1:1000–2000 dilution) in 0.5% BSA–PBS(+) (200 μl/well) for 1 h on ice (see Note 1). 5. Rinse the cells with ice-cold PBS(+) (1 ml/well) three times (see Note 2). 6. Bind HRP-conjugated anti-mouse IgG (1:1000–2000 dilution) in 0.5% BSA–PBS(+) (200 μl/well) for 1 h on ice.


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7. Rinse the cells with ice-cold PBS(+) (1 ml/well) 6–7 times (see Note 2). R Red reaction mix (200 μl/well) and incu8. Add Amplex bate on ice for 10–20 min in the dark (see Note 3).

9. Transfer 200 μl Amplex Red reaction mix from 24-well plates to black 96-well plates. R Red signal by a fluorescence plate 10. Read the Amplex reader (e.g., POLAR star Optima, BMG LABTECH) at 544 nm excitation and 590 nm emission wavelength.

11. Rinse the cells with 1 ml ice-cold PBS(+) once and add 70 μl RIPA buffer to lyse the cells for the BCA assay to measure the protein concentration (see Note 4). 12. Calculate the relative cell surface CFTR-3HA density based on the specific fluorescence signal. Normalize the fluorescence signal for the protein concentration of the respective well. Subtract the normalized fluorescence signal measured on non-transfected cells from the total fluorescence signal. Express the cell surface density of the CFTR variant as percentage of the wt-CFTR (Fig. 20.1b). 3.1.2. CFTR Internalization and Cell Surface Stability Measurement

Modification of the cell surface ELISA enables to determine the internalization rate and cell surface stability of CFTR-3HA (12, 20, 21). The CFTR-3HA internalization and the cell surface stability are measured by monitoring the disappearance of CFTRbound anti-HA antibody from the plasma membrane at 37◦ C. 1. Seed the cells as described in Section 3.1.1. Prepare multiple 24-well plates for different length of chase (e.g., time 0, 5, 60, and 120 min). 2. Bind the primary anti-HA antibody as described in Section 3.1.1. 3. Rinse the cells with ice-cold PBS(+) (1 ml/well) three times. 4. CFTR endocytosis is induced by the addition of the prewarmed (37◦ C) complete medium (+FBS). Incubate the cells for the desired time period to allow internalization (e.g., 2.5 and 5 min). Longer chase periods are required to measure the CFTR turnover at the plasma membrane at 37◦ C. Since CFTR appears to be less stable in HEK293 cells than in other non-polarized cells, 60- and 120-min incubation was chosen in this study. 5. After the incubation at 37◦ C, terminate the internalization by rinsing the cells with ice-cold PBS(+) (1 ml/well) two times (see Note 5). 6. Bind HRP-conjugated anti-mouse IgG as described in Section 3.1.1.

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7. Rinse the cells with ice-cold PBS(+) (1 ml/well) 6–7 times R Red signal as described in Secand measure the Amplex tion 3.1.1. 8. The CFTR internalization rate is expressed as the percentage of channel uptake after 5 min relative to the initial amount at cell surface (Fig. 20.1c). To determine the cell surface stability, the disappearance kinetics of cell surface labeled CFTR is plotted. The remaining amount of CFTR is expressed as the percentage of the initial amount. The channel turnover is indicated by the chase time that is necessary to decrease the CFTR cell surface density by 50% (Fig. 20.1d). 3.2. Methods for Detection of Native CFTR 3.2.1. Cell Culture and Lysis

1. The CFBE cells are maintained at 37◦ C in a humidified incubator containing 5% CO2 and allowed to grow until 90–95% confluent. At this point, using aseptic techniques, they are rinsed twice with HBS, rinsed with 5 ml PET, then covered with PET again, and returned to the incubator for 5 min. To the dislodged cells, 7 ml of media is added and the mixture placed in a 15-ml conical tube. Cells are pelleted gently at 10◦ C, at 800–1000 rpm in a Sorvall RT6000B refrigerated centrifuge, the media removed, cells dislodged by tapping, 4 ml of fresh media added, and 1 ml of the heterogeneous mixture added to each new T-75 flask containing 9 ml of medium. 2. Calu-3 cells are cultured in a similar manner with appropriate media and trypsin. Time for removal of these cells from plastic is longer, about 7 min. 3. Cells are lysed by rinsing the flask twice with PBS, then adding 1 ml of RIPA/PIT. Assays are performed to determine protein concentration.

3.2.2. Choice of Antigen

The predicted structure of CFTR suggests that there are six extracellular loops (27); however, four are less than five amino acids and are likely unsuitable for antibody production. The fourth loop has two consensus glycosylation sites for N-linked glycans which may mask antibody recognition of the epitope, rendering it an unsuitable candidate. The first loop has 15 amino acids:

GRIIASYDPDNKEER Several are polar, hydrophilic, and form an epitope likely accessible to antibodies. In order to restrict separation and hold the peptide into a loop, two cysteines were added at the carboxy and amino termini to form disulfide bonds and a third cysteine was added at the carboxy terminus for coupling.


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3.2.3. Synthesis of Antigen

The peptide is synthesized in solid phase on a Liberty Microwave Synthesizer (CEM Corporation, 3100 Smith Farm Road, Mathews, NC 28106) using an FMOC synthesis protocol (University of Pittsburgh Peptide Synthesis Facility). Briefly, synthesis is performed by stepwise addition of activated amino acids to the solid support (preloaded Wang resin) starting at the carboxy and proceeding to the amino terminus. Activation of amino acids is performed by DIPEA/HOBT/TBTU chemistry. For regioselective cyclization between cysteine positions 1 and 17 (carboxy to amino), trityl-protecting groups are used and the N-terminal cysteine is protected by a PMeOBZl group. At the end of synthesis, the peptide is cleaved from the resin by reagent R (90% TFA, 5% thioanisole, 3% ethanedithiol, and 2% anisole) and subjected to multiple ether extractions. The crude peptide is analyzed, characterized, and purified by gel filtration (G-25 column) and reversed-phase, high-performance liquid chromatography (RPHPLC, 486 and 600E by Waters Corporation), and later the correct mass is confirmed by MALDI-TOF mass spectroscopy (The Voyager-DE STR Biospectrometry Workstation). Cyclization of purified peptide is performed by a hydrogen peroxide method and confirmed by Elman’s test. At the end, the PMeOBZL group is removed from the N-terminal cysteine and the final product is re-purified by HPLC and confirmed by mass spectrometry.

3.2.4. Coupling of Peptide to Keyhole Limpet Hemocyanin

In order for the peptide hapten to elicit an adequate immune response in the rabbit, it is coupled with KLH, a large, foreign protein, following manufacturer’s directions (Pierce): 1. Ten milligrams of activated carrier is reconstituted in 1 ml dH2 O and 10 mg of hapten is added. This carrier/peptide solution is allowed to react for 2 h at room temperature. R dialysis cassette 2. The solution is dialyzed in a Slide-A-Lyzer ◦ (Pierce), 10,000 MWCO, at 4 C overnight, removed from cassette, aliquoted, and frozen.

3.2.5. Production of Antibodies in Rabbits

Before any treatment is commenced in rabbits, a pre-immune bleed (about 20 ml) is removed from the medial ear artery with a 21-gauge butterfly needle and the serum is harvested for use as negative controls in future experiments. To carry out this procedure, blood is allowed to clot for 1 h at room temperature, separated from the sides of the tube with the wooden end of a long, cotton-tipped applicator, then spun at 10,000×g for 10 min. The serum (supernatant) is recovered from the pellet (red blood cells), aliquoted in quantities that will not necessitate repeated freeze/thaws, and frozen at –80◦ C. To prepare for injections of antigen, an emulsion is made with equal volumes of coupled peptide (500 μg per rabbit) in PBS and R Freund’s complete adjuvant (Pierce) adjuvant. On day 1, Imject is used for the first series of 5–7 subcutaneous injections and on

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3.2.6. Enzyme-Linked Immunosorbent Assay (ELISA)

In preparation for this experiment, (a) allow diethanolamine buffer to come to room temperature and (b) make appropriate dilutions of antiserum into TBSTB in microcentrifuge tubes, e.g., start with 1:100, then proceed with progressive twofold dilutions in the remaining 11 tubes. 1. Coat 96-well microtiter plate with antigen by placing 200 μl carbonate buffer to each well and adding 50 pmol antigen to each well. Cover plate with parafilm and incubate at room temperature overnight. 2. Block the plate. Fill each well by adding TBSTB. Incubate at 37◦ C for 1 h. 3. Wash plate three times using TBST. 4. Add 100 μl antiserum into each well. Incubate at 37◦ C for 1 h covered with parafilm. 5. Wash plate three times with TBST. 6. Make a 1:5000 dilution of secondary Ab–alkaline phosphatase conjugate (goat anti-rabbit) in TBSTB and add 100 μl to each well. Incubate for 2 h at 37◦ C. 7. Wash plate three times using TBST. 8. Dissolve para-nitrophenylphosphate tablets (one 5 mg tablet/5 ml diethanolamine buffer). Add 100 μl of this substrate to each well. Incubate for 30 min at room temperature protected from light. 9. Read plate at 414 nm subtracting out nonspecific background of the plate at 650 nm. 10. Binding curves are defined at half maximal values with 50 pmol of antigen (Fig. 20.2a).

3.2.7. Immunoprecipitation

Immunoprecipitations are carried out as described by Harlow and Lane with modifications: 1. Rinse flasks twice with PBS, then remove PBS. 2. Add RIPA, scrape cells, and place lysate in a microcentrifuge tube. 3. Sonicate lysates, then rotate for 1 h at 4◦ C. 4. Centrifuge for 3 min at 13,000×g. Recover the supernatant and perform a protein assay. 5. Place 500 μg of soluble lysate in a microcentrifuge tube. Add 3 μl ECL1 antibody and rotate overnight at 4◦ C. 6. Add 50 μl protein bead slurry and rotate for 7 h or overnight.


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A

B

kDa

1

2

3

4

C

Fig. 20.2. (a) Data from a quantitative ELISA for antibody recognition of the first extracellular loop of CFTR, ECL1. Fifty picomoles of ECL1 peptide was used with a repetitive series of 1:2 dilutions of the antiserum. Binding curves were half-maximal at 50 pM, at an antibody dilution of 1:100,000. (b) The ECL1 antibody effectively immunoprecipitates CFTR from CFBE cells. Lanes: 1, parental CFBE; 2, CFBE-WT; 3, CFBE-F508del; 4, CFBE-F508del following 48 h incubation at 27◦ C. Although effective for immunoprecipitation, the ECL1 antibody does not recognize denatured CFTR by immunoblot (data not shown). (c) CFTR detection by immunofluorescence in polarized Calu-3 cells. Left, peptide competition of ECL1 antibodies with synthetic peptide prior to use. Right, ECL1 staining of non-permeabilized Calu-3 monolayer using the IgG fraction. Cells cultured on Transwell filter for 1 week were imaged by confocal microscopy at the first detectable apical signal, which creates the patchy fluorescence patter due to the uneven cell layer.

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7. Centrifuge immunocomplexes and wash pellets times with 1 ml RIPA. 8. Remove RIPA, add sample buffer, and incubate at 40◦ C for 10 min. 9. Load samples onto a 6% gel for SDS-PAGE. 3.2.8. Immunoblot

1. Block membranes, into which proteins have been transferred, in TBSTM for 1 h at room temperature. 2. Incubate membrane with primary antibody at appropriate dilution (e.g., 217 monoclonal 1:5000) for 1 h at room temperature. 3. Wash three times for 5 min each in TBST. 4. Incubate membrane in appropriate peroxidase-conjugated secondary antibody at a dilution of 1:20,000 for 2 h at room temperature. 5. Wash three times for 5 min each. 6. Mix substrate chemiluminescent detection reagents one and two (GE Healthcare) in equal amounts and apply to toweldried membranes for 1 min. 7. Blot membranes dry and expose to film for optimal exposure (Fig. 20.2b).

3.2.9. Immunofluorescence

1. Seed Calu-3 cells onto filters (Snapwells with polycarbonate membrane) (Costar) and maintain until confluent. 2. Wash filters in PBS. 3. Add agonist for 10 min at room temperature. 4. From this point on, perform all steps with ice-cold buffers and keep filters on ice. 5. Wash filters three times with PBS, followed by three washes with PBSBG. 6. Block non-specific binding with PBSBG-GS for 1 h. 7. Wash three times with ice-cold PBSBG. 8. Incubate cells with primary antibody diluted 1:200 in PBSBG for 1 h. 9. Wash three times with PBSBG. 10. Incubate cells with secondary antibody, e.g., goat antirabbit CY3 1:1000 for 1 h. 11. Wash three times with PBSBG followed by three washes in PBS. 12. Cut filters from Snapwells and orient them upside down onto coverslips. Place Gelvatol onto a microscope slide, position over the filter, and press (Fig. 20.2c).


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4. Notes 4.1. Detection of HA-CFTR

1. It is recommended to optimize the antibody concentration in order to maximize the specific signal. This could be achieved by using serial dilutions of the primary and secondary antibodies on transfected and non-transfected cells in pilot studies. Incubation should be done at 4◦ C to prevent the CFTR–antibody internalization. 2. Add the solution carefully by pipetting down the sides of the wells to avoid detachment of cells. Avoid drying the cells during the washing by immediately adding the medium after aspiration. 3. If the fluorescence signal is weak, extend the incubaR tion time. If you have too strong signal (e.g., Amplex Red becomes pink in 1–2 min), you need to dilute the R Red to avoid the signal saturation. antibodies/Amplex 4. This is not necessary if the cell number in each sample is same. If the cell number is different between samples, the fluorescence signal normalization is required for protein concentration, which reflects the cell number. 5. During the 37◦ C incubation, keep the other plates on ice to prevent the CFTR–antibody complex internalization.

Acknowledgments Resources providing support for this work in the Frizzell lab include grants from the NIH (DK068196 and DK 072506) and the Cystic Fibrosis Foundation (CFF R883-CR07 and FRIZZE05XX0). Experimental work in the laboratory of Gergely Lukacs was funded by the NIH, Cystic Fibrosis Folding Consortium, CIHR, and CFI. Tsukasa Okiyoneda was supported by a postdoctoral fellowship from the Canadian Cystic Fibrosis Foundation. References 1. Cheng, S. H., Gregory, R. J., Marshall, J., Paul, S., Souza, D. W., White, G. A., et al. (1990) Defective intracellular transport and processing of CFTR is the molecular basis of most cystic fibrosis. Cell 63, 827–834. 2. Zhang, F., Kartner, N., and Lukacs, G. L. (1998) Limited proteolysis as a probe for

arrested conformational maturation of delta F508 CFTR. Nat Struct Biol 5, 180–183. 3. Du, K., Sharma, M., and Lukacs, G. L. (2005) The DeltaF508 cystic fibrosis mutation impairs domain-domain interactions and arrests post-translational folding of CFTR. Nat Struct Mol Biol 12, 17–25.

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4. Denning, G. M., Anderson, M. P., Amara, J. F., Marshall, J., Smith, A. E., and Welsh, M. J. (1992) Processing of mutant cystic fibrosis transmembrane conductance regulator is temperature-sensitive. Nature 358, 761–764. 5. Welch, W. J. (2004) Role of quality control pathways in human diseases involving protein misfolding. Semin Cell Dev Biol 15, 31–38. 6. Wang, X., Venable, J., LaPointe, P., Hutt, D. M., Koulov, A. V., Coppinger, J., et al. (2006) Hsp90 cochaperone Aha1 downregulation rescues misfolding of CFTR in cystic fibrosis. Cell 127, 803–815. 7. Zhang, Y., Nijbroek, G., Sullivan, M. L., McCracken, A. A., Watkins, S. C., Michaelis, S., et al. (2001) Hsp70 molecular chaperone facilitates endoplasmic reticulum-associated protein degradation of cystic fibrosis transmembrane conductance regulator in yeast. Mol Biol Cell 12, 1303–1314. 8. Sun, F., Mi, Z., Condliffe, S. B., Bertrand, C. A., Gong, X., Lu, X., et al. (2008) Chaperone displacement from mutant cystic fibrosis transmembrane conductance regulator restores its function in human airway epithelia. FASEB J 22, 3255–3263. 9. DeCarvalho, A. C., Gansheroff, L. J., and Teem, J. L. (2002) Mutations in the nucleotide binding domain 1 signature motif region rescue processing and functional defects of cystic fibrosis transmembrane conductance regulator delta F508. J Biol Chem 277, 35896–35905. 10. Teem, J. L., Carson, M. R., and Welsh, M. J. (1996) Mutation of R555 in CFTR-delta F508 enhances function and partially corrects defective processing. Receptors Channels 4, 63–72. 11. Chang, X. B., Cui, L., Hou, Y. X., Jensen, T. J., Aleksandrov, A. A., Mengos, A., et al. (1999) Removal of multiple arginine-framed trafficking signals overcomes misprocessing of delta F508 CFTR present in most patients with cystic fibrosis. Mol Cell 4, 137–142. 12. Pedemonte, N., Lukacs, G. L., Du, K., Caci, E., Zegarra-Moran, O., Galietta, L. J., et al. (2005) Small-molecule correctors of defective DeltaF508-CFTR cellular processing identified by high-throughput screening. J Clin Invest 115, 2564–2571. 13. Van Goor, F., Straley, K. S., Cao, D., Gonzalez, J., Hadida, S., Hazlewood, A., et al. (2006) Rescue of DeltaF508-CFTR trafficking and gating in human cystic fibrosis airway primary cultures by small molecules. Am J Physiol Lung Cell Mol Physiol 290, L1117–L1130.

14. Pyle, L. C., Balch, W. E., Lukacs, G., Braakman, I., Guggino, W. B., Thomas, P. J., et al. (2010) Developing a cellular road map for correctors of protein misfolding: a consortium approach. Nat Rev Drug Disc. (submitted). 15. Lukacs, G. L., Segal, G., Kartner, N., Grinstein, S., and Zhang, F. (1997) Constitutive internalization of cystic fibrosis transmembrane conductance regulator occurs via clathrin-dependent endocytosis and is regulated by protein phosphorylation. Biochem J 328 (Pt 2), 353–361. 16. Prince, L. S., Peter, K., Hatton, S. R., Zaliauskiene, L., Cotlin, L. F., Clancy, J. P., et al. (1999) Efficient endocytosis of the cystic fibrosis transmembrane conductance regulator requires a tyrosine-based signal. J Biol Chem 274, 3602–3609. 17. Benharouga, M., Haardt, M., Kartner, N., and Lukacs, G. L. (2001) COOH-terminal truncations promote proteasome-dependent degradation of mature cystic fibrosis transmembrane conductance regulator from postGolgi compartments. J Cell Biol 153, 957– 970. 18. Denning, G. M., Ostedgaard, L. S., Cheng, S. H., Smith, A. E., and Welsh, M. J. (1992) Localization of cystic fibrosis transmembrane conductance regulator in chloride secretory epithelia. J Clin Invest 89, 339–349. 19. Denning, G. M., Ostedgaard, L. S., and Welsh, M. J. (1992) Abnormal localization of cystic fibrosis transmembrane conductance regulator in primary cultures of cystic fibrosis airway epithelia. J Cell Biol 118, 551–559. 20. Sharma, M., Pampinella, F., Nemes, C., Benharouga, M., So, J., Du, K., et al. (2004) Misfolding diverts CFTR from recycling to degradation: quality control at early endosomes. J Cell Biol 164, 923–933. 21. Glozman, R., Okiyoneda, T., Mulvihill, C. M., Rini, J. M., Barriere, H., and Lukacs, G. L. (2009) N-glycans are direct determinants of CFTR folding and stability in secretory and endocytic membrane traffic. J Cell Biol 184, 847–862. 22. Barriere, H., Bagdany, M., Bossard, F., Okiyoneda, T., Wojewodka, G., Gruenert, D., et al. (2009) Revisiting the role of cystic fibrosis transmembrane conductance regulator and counterion permeability in the pH regulation of endocytic organelles. Mol Biol Cell 20, 3125–3141. 23. Robert, R., Carlile, G. W., Pavel, C., Liu, N., Anjos, S. M., Liao, J., et al. (2008) Structural analog of sildenafil identified as a novel corrector of the F508del-

CFTR Folding Consortium Reagents and Protocols CFTR trafficking defect. Mol Pharm 73, 478–489. 24. Carlile, G. W., Robert, R., Zhang, D., Teske, K. A., Luo, Y., Hanrahan, J. W., et al. (2007) Correctors of protein trafficking defects identified by a novel high-throughput screening assay. ChemBioChem 8, 1012–1020. 25. Voller, A., Bidwell, D., Rose, N. R., Friedman, H., and Fahey, J. L. (1986) Enzymelinked immunosorbent assay, in Manual of Clinical Laboratory Immunology, vol. 3.

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American Society of Microbiology, Washington, DC, pp. 99–109. 26. Harlow, E., and Lane, D. (1988) Immunoprecipitation, in Antibodies: A Practical Approach. 27. Riordan, J. R., Rommens, J. M., Kerem, B., Alon, N., Rozmahel, R., Grzelczak, Z., et al. (1989) Identification of the cystic fibrosis gene: cloning and characterization of complementary DNA. Science 245, 1066–1073.

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Chapter 21 Evaluation of the Disease Liability of CFTR Variants Patrick R. Sosnay, Carlo Castellani, Mary Corey, Ruslan Dorfman, Julian Zielenski, Rachel Karchin, Christopher M. Penland, and Garry R. Cutting Abstract Over 1600 novel sequence variants in the CFTR gene have been reported to the CF Mutation Database (http://www.genet.sickkids.on.ca/cftr/Home.html). While about 25 mutations are well characterized by clinical studies and functional assays, the disease liability of most of the remaining mutations is either unclear or unknown. This gap in knowledge has implications for diagnosis, therapy selection, and counseling for patients and families carrying an uncharacterized CFTR mutation. This chapter will describe a critical approach to assessing the disease implications of CFTR mutations utilizing clinical data, literature review, functional testing, and bioinformatic in silico methods. Key words: CFTR mutations, CFTR variants, CFTR polymorphisms, mutation prediction algorithms, in silico testing, bioinformatic analysis, genotype–phenotype relationship.

1. Introduction Defining disease liability: Disease liability in reference to a germline mutation refers to the capability of that allele to influence a given disease phenotype. For example, an individual with the p.Glu6Val (HbS) mutation in each beta-globin gene is expected in all cases to have sickle-cell anemia. Likewise, in the case of cystic fibrosis (CF), the full penetrance of the most common deleterious allele in the cystic fibrosis transmembrane conductance regulator gene (CFTR) (p.Phe508del, c.1521_1523delCTT) is well documented. However, unlike sickle cell anemia there are over 1600 variants in CFTR, many M.D. Amaral, K. Kunzelmann (eds.), Cystic Fibrosis, Methods in Molecular Biology 742, DOI 10.1007/978-1-61779-120-8_21, © Springer Science+Business Media, LLC 2011

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of which have been reported in one or only a few individuals. The 23 alleles included in the American College of Medical Genetics (ACMG) panel on screening for CF and a few other mutations have been identified as disease causing by evaluating patients carrying one of these mutations and p.Phe508del on the other chromosome and by functional testing (1, 2). However, the remaining CFTR variants identified in the CF Mutation Database (CFMD) have not been rigorously examined. Systematic and objective evaluation of the disease-causing potential of CFTR mutations is important as genetic testing for CFTR mutations is being performed more frequently for newborn screening, carrier screening, and CF diagnosis and for consideration of mutation-specific therapeutics (3). The lessons and experience learned from developing an algorithm to evaluate CFTR mutations should be relevant to complex diseases with a heritable component (4, 5). Challenges created by multiple CFTR variants: Individuals with CF have mutations in both copies of their CFTR gene, as expected for a recessive Mendelian disorder. The Hardy– Weinberg law establishes the relationship between the allele frequency and the genotype frequency in a population (Fig. 21.1). The most common CFTR mutation p.Phe508del accounts for 70% of CF alleles and has a frequency of approximately 2.5% in the general population (6, 7). Both of these frequencies assume a population of Caucasian European descent, though there may be remarkable differences among distinct geographical areas (7, 8). From the frequency of p.Phe508del in CF patients, one can estimate that 49% of individuals with CF have two copies of p.Phe508del allele (p2 = (0.7)2 = 0.49). The remaining individuals will have either one p.Phe508del with another CFTR mutation (2pq = 2 × 0.7 × 0.3 = 0.42) or two non-p.Phe508del alleles (q2 = (0.3)2 = 0.09). When genotyping with the ACMG panel (which accounts for 85% of the CF alleles) one can predict that 27% of individuals with CF will carry at least one non-ACMG CF mutation (2 × 0.85 × 0.15 + (0.15)2 ). If enough CFTR mutations are well characterized to account for 95% of CF alleles, then only 10% of CF patients would have an uncharacterized CFTR

Fig. 21.1. The Hardy–Weinberg equation for population genetics. CFTR has many potential variants. The frequencies that appear in the text use this equation to compare genotypes when different percentages of the alleles are characterized.

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mutation (9). The effect of increasing the fraction of CF alleles known to cause disease upon the fraction of patients with fully informative genotypes is illustrated in Fig. 21.2. Another way of looking at this challenge is demonstrated in Fig. 21.3. Mutations such as p.Phe508del and p.Met470Val occur commonly, and therefore the confidence in determination of the effect of each mutation is high. Rare mutations constitute not only a large percentage of the total variation in CFTR but also a small fraction of all CF alleles. These uncharacterized mutations are infrequently evaluated and therefore there is low confidence in any prediction of disease liability. Although we are providing population-based estimates here, the difficulty with uncharacterized CFTR mutations is acute in individual cases. For example, consider the challenge of an uncharacterized CFTR mutation in an asymptomatic child detected by a newborn screen. There are uncertain ramifications of diagnosis and a lifetime of costly therapies to consider. Determining the Effect of Variants on CFTR Function: It is reasonable to predict that each CFTR variant alters protein function in some manner along a spectrum from null (no function) to

Fig. 21.2. Percentage of CF total mutations, total alleles in CF patients, and individuals with genotypes that would be characterized based on the number of mutations evaluated. The fraction of total mutations is based on 1600 CFTR variants described in the CFMD. The percentage of total alleles is based on the worldwide CFTR2 database (Cutting, unpublished results). The percentage of individuals in which both CFTR mutations are identified uses the allele percentage and the Hardy–Weinberg law.

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Fig. 21.3. The relationship between CFTR variant disease liability and (a) the relative number of patients that carry that variant; (b) the number of variants expressed as the percentage of total variants in CFTR; (c) the confidence of our ability to determine if a variant is disease causing or neutral. Common variants or polymorphisms as well as mutations that are often seen in CF patients are frequently seen and therefore one can determine with confidence their disease liability. Infrequently seen mutations affect few patients, but these represent a large percentage of the total CFTR variation described. Because these mutations are seen less there are less rigorous evaluations and therefore the confidence about the determination of their disease liability is less.

near-wild type or even, theoretically, enhanced protein function. Disease-causing mutations (such as p.Phe508del) are expected to cause CFTR to be dysfunctional. At the other end of the spectrum are amino acid substitutions (such as p.Met470Val) that cause only minor alterations in function and are not associated with disease (10–12). For CF diagnostic purposes, one would ideally have all CFTR variants characterized as either a disease-causing mutation (like p.Phe508del) or a neutral (like p.Met470Val). The key challenge is to identify the threshold at which a reduction in protein function is enough to cause disease. Features of individuals carrying a given variant, functional evaluation of the variant in cell-based assays, or predictive evaluation of the consequences of the variant each may be used to evaluate where along the spectrum between deleterious and near wild type a given variant belongs. Correlating CFTR Genotype with CF Phenotype: The severity spectrum and threshold for CFTR differ across organ systems, illustrated in Fig. 21.4 (13). This is a result of the differing role of CFTR in pathogenesis for each of these organ systems. CF carriers may demonstrate single organ system traits seen in CF,


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Fig. 21.4. Genotype and specific organs system phenotype correlation. The relative penetrance of separate organ systems is shown relative to the degree of CFTR dysfunction. CF carriers, with only one CF causing CFTR mutation, may have some of the traits seen in CF patients. In these cases, CFTR is acting as a modifier gene interacting with other genetic and environmental predispositions, rather than the sole etiology of the organ dysfunction (ABPA = allergic bronchopulmonary aspergillosis).

but with much lower penetrance than individuals with two CFTR mutations (14–16). The male reproductive tract is the most sensitive tissue to detect CFTR dysfunction. Individuals who carry a single CFTR mutation, as well as individuals with mutations in each CFTR gene that cause only mild reductions in function, may have obstructive azoospermia without any other signs of disease (13, 14, 17, 18). Pancreatic and lung tissue require more severe disruption of CFTR function to manifest features of CF. CFTR mutations have been arranged into five functional classes by Welsh and Smith according to the defect in protein function (19). Classes 1–3 CFTR mutations which cause the most severe disruption of CFTR function are associated with pancreatic insufficient disease compared to classes 4 and 5 CFTR mutations which lead to milder impairment in CFTR function and are associated with partially retained pancreatic function (20). This categorization of CFTR mutations by their effect on pancreatic function is problematic and will be discussed below. Additionally, how well correlated the severity of organ system pathology is

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with the degree of CFTR dysfunction is different across different organ systems, is subject to different genetic and environmental modifiers for each organ system, and varies with time. These complicating factors confound researchers’ and clinicians’ ability to predict the degree of lung dysfunction that will be associated with a CFTR genotype.

2. Tools to Evaluate CFTR Variants for Disease Liability

Clinical Data: One method to assess pathogenicity is to view every CFTR mutation occurring in an individual meeting clinical diagnostic criterion for CF as disease causing. However, there are a few caveats to this approach. First, although we are attempting to assess the disease liability for the mutation, we are evaluating the mutation in the context of the recessive phenotype caused by alterations in both CFTR genes. Heterozygote carriers do not manifest CF (although they may have a predisposition to other conditions). Therefore to create a CF phenotype both copies of CFTR must be dysfunctional. It is important therefore when evaluating the disease liability of a CFTR mutation of unknown effect to analyze patients carrying a known CF-causing mutation in their other CFTR gene. The severity and commonness of p.Phe508del facilitate its use as a consistent background on which to compare novel mutations (21–23). Second, discerning the disease liability of a mutation when multiple mutations exist in the same CFTR gene can be a substantial challenge. These “complex alleles” may harbor covert point mutations in the coding or noncoding regions of the gene or a large insertion, deletion, or duplication (24, 25). For example, the variant p.Ile148Thr was initially felt to be disease causing, but further examination found that the mutation occurred at a higher rate in healthy individuals undergoing carrier screening than in CF patients (26). Subsequent studies revealed that most CF patients with p.Ile148Thr also carry the two amino acid deletion c.3067_3072del (27). The latter mutation in isolation causes CF whereas p.Ile148Thr in isolation does not. Other CFTR variants may not individually cause disease, but can cause disease when they occur in a CFTR gene with other variants. For example, the disease liability of the ACMG mutation p.Arg117His is dependent on polythymidine variants in the flanking exon (28). When this mutation is associated with the “5T” polythymidine tract, p.Arg117His has a higher penetrance for CF, whereas longer polythymidine tracts (“7T” or “9T”) are associated with obstructive azoospermia or no disease at all. For these reasons it is critical to confirm that a mutation in isolation is deleterious both by demonstrating the clinical


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traits consistent with CF in several individuals that carry the novel mutation and by demonstrating its dysfunction on a cellular level. As an additional test, variants identified in healthy carriers with a CF-causing CFTR mutation on the other chromosome must be neutral. Third, direct measures of CFTR function (sweat chloride and nasal potential differences (NPD) measurements) are not highly correlated with the morbidity and mortality of the disease. Other traits, such as lung dysfunction and airway bacterial colonization, more useful in predicting clinical outcome in CF, are usually normal at birth and only manifest over time. Furthermore, these traits show high individual variability and are influenced by genetic and environmental modifiers (29). With these limitations in mind, the effect of a CFTR mutation on an individual’s phenotype is the single most important measure of the medical relevance of that mutation. The specific features used to clinically evaluate CFTR function are described below.

3. Direct Assays of In Vivo CFTR Function

Sweat Chloride Measurement: The determination of ion concentration in sweat induced by pilocarpine administration is an objective method to differentiate CF from other diseases of the lungs or pancreas. The procedure is standardized, accurate, and highly reproducible (30, 31). In the context of a positive newborn screen or clinical symptoms of CF, individuals with a sweat chloride concentration higher than 60 mEq/L have two CF-causing CFTR mutations. This is a specific threshold as lower sweat values do not exclude two disease-causing mutations (32). However, only 3.5% of CF patients in the USA had sweat chloride concentration less than 60 mEq/L and only 1.2% had values less than 40 mEq/L (33). The thresholds for sweat chloride measurement were recently changed (>60 mEq/L: positive; 30–60 mEq/L for infants, 40–60 mEq/L for children over 6 months: intermediate; and

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